Method Article

Real-Time Void Spot Assay

DOI:

10.3791/64621

⸱

February 10th, 2023

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

In This Article

Summary

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This article describes a new method to study mouse voiding behavior by incorporating video monitoring in the conventional void spot assay. This approach provides temporal, spatial, and volumetric information on the voiding events and details of mouse behavior during the light and dark phases of the day.

Abstract

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Normal voiding behavior is the result of the coordinated function of the bladder, the urethra, and the urethral sphincters under the proper control of the nervous system. To study voluntary voiding behavior in mouse models, researchers have developed the void spot assay (VSA), a method that measures the number and area of urine spots deposited on a filter paper lining the floor of an animal’s cage. Although technically simple and inexpensive, this assay has limitations when used as an end-point assay, including a lack of temporal resolution of voiding events and difficulties quantifying overlapping urine spots. To overcome these limitations, we developed a video-monitored VSA, which we call real-time VSA (RT-VSA), and which allows us to determine voiding frequency, assess voided volume and voiding patterns, and make measurements over 6 h time windows during both the dark and light phases of the day. The method described in this report can be applied to a wide variety of mouse-based studies that explore the physiological and neurobehavioral aspects of voluntary micturition in health and disease states.

Introduction

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Urine storage and micturition are coordinated by a central circuitry (central nervous system) that receives information about the bladder filling status through the pelvic and hypogastric nerves. The urothelium, the epithelium that lines the urinary tract from the renal pelvis to the proximal urethra, forms a tight barrier to the metabolic waste products and pathogens present in urine. It is an integral component of a sensory web, which senses and communicates the filling state of the bladder to underlying tissues and afferent nerves1,2. Disruption of the urothelial barrier, or alterations in urothelial mechanotransduction pathways, can lead to voiding dysfunction along with lower urinary tract symptoms such as frequency, urgency, nocturia, and incontinence3,4,5,6,7. Likewise, aging, diabetes, lower urinary tract infections, interstitial cystitis, and other disease processes that affect the urinary bladder, or the associated circuitry that controls its function, are known to cause bladder dysfunction8,9,10,11,12,13,14,15,16,17,18,19. A better understanding of normal and abnormal voiding behavior depends on the development of methods that can reliably discriminate among different urination patterns. 

Traditionally, the voluntary voiding behavior of mice has been studied using the void spot assay (VSA), developed by Desjardins and colleagues20, and broadly adopted due to its simplicity, low cost, and noninvasive approach8,21,22,23,24. This assay is typically performed as an endpoint assay, in which a mouse spends a defined amount of time in a cage lined by a filter paper, which is subsequently analyzed by counting the number and assessing the size of urine spots when the filter paper is placed under ultraviolet (UV) light (the urine spots fluoresce under these conditions)20. Despite these many advantages, the traditional VSA presents some major limitations. Because mice often urinate in the same areas, investigators have to restrict the duration of the assay to a relatively short period of time (≤4 h)25. Even when the VSA is performed over shorter time periods, it is almost impossible to resolve small void spots (SVSs) that fall over large void spots or, to discriminate SVSs from the carryover of urine adhered to tails or paws. It is also very difficult to distinguish if SVSs are a consequence of frequent but individual voiding events (a phenotype that is often observed in response to cystitis4,26), or due to post-micturition dribbling (a phenotype associated with bladder outlet obstruction27). Furthermore, the desire to complete the assay during working hours, coupled with difficulties accessing housing facilities when the lights are turned off, often limits these assays to the light period of the 24 h circadian cycle. Thus, these time constraints prevent the evaluation of mouse voiding behavior during their active night phase, lessening the ability to analyze specific genes or treatments that are governed by circadian rhythms. 

To overcome some of these limitations, researchers have developed alternative methods to assess voiding behavior in real time26,28,29,30,31,32. Some of these approaches involve the use of expensive equipment such as metabolic cages26,28,29, or the use of thermal cameras30; however, these too have limitations. For example, in metabolic cages, urine tends to adhere to the wires of the mesh floor and to the walls of the funnel, reducing the amount of urine that is collected and measured. Thus, it can be difficult to accurately collect data about small voids. Moreover, metabolic cages do not provide information about the spatial distribution of the voiding events (i.e., urination in the corners vs. the center of the chamber). Given that long-wavelength infrared radiation used by the thermographic cameras does not penetrate solids, voiding activity assessed by video thermography must be performed in an open system, which can be challenging with active mice, as they can jump several inches in the air. Another system is the automated voided stain on paper (aVSOP) approach33, which consists of rolled filter paper that winds up at a constant speed below the wire mesh floor of a mouse cage. This approach prevents paper damage and the overlap of urine spots that occur in the classical VSA, and its implementation allows the investigator to perform experiments over several days. However, it does not provide the investigator with precise timing of the voiding events, and there is no ability to examine behavior and how it correlates with spotting. To obtain this information, researchers have incorporated video-monitoring to voiding assays, an approach that allows the simultaneous assessment of mouse activity and urination events31,32. One approach consists of placing a blue light emitting diode (LED) and a video-camera with a green fluorescence protein filter set under the experimental cage to visualize the voiding events, and an infrared LED and a video-camera above the cage to capture mouse position32. This setup has been used to monitor voiding behavior while performing fiber photometry; however, the brightly lit environment of this system required the investigators to treat their mice with a diuretic agent to stimulate voiding. In another experimental design, wide-angle cameras were placed above and below the experimental cage to visualize mouse motor activity and urination events, respectively. In this case, urine spots deposited on a filter paper lining the cage’s floor were revealed by illuminating the filter paper with UV lights placed under the cage31. This setup was used in short assays, 4 min in duration, during the light phase of the day to study the brainstem neurons involved in voluntary voiding behavior31. The suitability of this system for its use during the dark phase or for periods of time >4 min was not reported. 

In this article, a method is described that enhances the traditional VSA by allowing for long-term video monitoring of mouse voiding behavior. This cost-effective approach provides temporal, spatial, and volumetric information about voiding events for extended periods of time during the light and dark phases of the day, along with details related to mouse behavior3,4,34. Detailed information for the construction of the voiding chambers, the implementation of a real-time VSA (RT-VSA), and the analysis of the data is provided. The RT-VSA is valuable for researchers seeking to understand the physiological mechanisms that control the function of the urinary system, to develop pharmacological approaches to control micturition, and to define the molecular basis of disease processes that affect the lower urinary tract. 

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Protocol

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Urothelial Piezo1/2 double knockout mice (Pz1/2-KO, genotype: Piezo1fl/fl;Piezo2fl/fl;Upk2CRE+/-) and controls (Pz1/2-C, genotype: Piezo1fl/fl; Piezo2fl/fl; Upk2CRE-/-) were generated in-house from parental strains obtained from the Jax laboratories (Piezo1fl/fl strain # 029213; Piezo2fl/fl strain # 027720; Upk2CRE+/- strain # 029281). Both female (1.5–3 months old and 17–20 g in weight) and male (2–4 months old and 23–29 g in weight) mice were used in the experiments. For cyclophosphamide-induced cystitis experiments, wild-type C57Bl/6J females (3 months old and ~20 g in weight) were used (the Jackson laboratories, strain # 000664). Animals were housed and the experiments were performed at the University of Pittsburgh Animal Care Facility under the approval of the University of Pittsburgh Institutional Animal Care and Use Committee. All animal experiments were performed in accordance with relevant guidelines and regulations of the Public Health Service Policy on Humane Care and Use of Laboratory Animals and the Animal Welfare Act.

1. Assembly of cages for real-time void spot assay (RT-VSA)

  1. The RT-VSA rig consists of an UV stand that holds two UV light bulbs and two wide-angle cameras (bottom cameras), which are used to record voiding activity during the light phase of the day. Arrange two mouse chambers to rest on the stand. Attach wide angle cameras (top cameras) to the lid of each mouse chamber, used to record voiding activity during the dark phase of the day (Figure 1A).
  2. Construct the frame of the UV stand and mouse chambers from 1 in x 1 in T-slotted aluminum profiles. See Table 1 for a list of the components used to build two mouse chambers and the UV stand and Figure 1B–D for the different views of the assembled components and dimensions.
  3. Construct each mouse chamber with eight T-slotted aluminum profiles cut to 10 in and four cut to 14.75 in. Start by assembling the bottom of the mouse chamber and use standard end fasteners (Table 1) to assemble the T-slotted profiles according to Figure 1. Mount the 38.5 cm x 26.5 cm UV transmitting acrylic in the internal channel of the profiles to build the floor of the mouse chambers.
    NOTE: For detailed information on how to assemble the T-slotted profiles with standard end fasteners, visit the company webpage.
  4. Build the walls of the mouse chamber with the rest of the profiles. Mount the 38.5 cm x 21.5 cm and 26.5 cm x 21.5 cm abrasion resistant (AR) polycarbonate panels in the profiles to assemble the external walls of the mouse chamber. Use the 37.5 cm x 23.9 cm and 24.4 cm x 23.9 cm AR polycarbonate panels to build the interior of the mouse chamber.
  5. Secure the AR polycarbonate panels with standard end fastener 1/4-20 tread. See instructions of how to mount the panels in the profiles on the webpage in the NOTE of step 1.3. Use commercial silicone caulk to seal the junctions between the internal panels.
  6. Use two 12 in and two 14.75 in T-slotted profiles to assemble the lid of the cage as indicated in Figure 1B. Mount a 38.5 cm x 26.5 cm AR polycarbonate panel in the internal channel of the profiles to complete the lid.
  7. Mount the 12 in camera profile support perpendicular to the long axes of the top of the cage and secure with two lite transition inside corner brackets (marked 3 in Figure 1B). Use a straight flat plate (marked 2) as the webcam mounting support and mount it as shown in Figure 1B. Attach the camera to the support with the camera mounting screw.
  8. Use the 10 series standard lift-off hinge right hand assembly to attach the lid to the body of the mouse chamber (marked 1 in Figure 1B).
  9. Construct the UV stand with four T-slotted profiles cut to 40 in, four T-slotted profiles cut to 32 in, and four T-slotted profiles cut to 10 in, according to Figure 1C,D. Mount a 82.5 cm x 26.5 cm acrylic mirror sheet in the profiles to build the bottom of UV chamber.
  10. Use the 82.5 cm x 30.5 cm and 26.5 cm x 30.5 cm acrylic mirror sheet to construct the wall of the UV stand.
  11. Attach the 10 series five-hole T flat plates (marked 2 in Figure 1C), the 10 series five-hole L flat plates (marked 3 in Figure 1C), and the 10 series five-hole tee flat plates (marked 1 in Figure 1C) to secure the UV stand.
  12. Mount the bottom cameras in a T-slotted profile cut to 32 in and affix to the bottom of the stand with two lite transition corner brackets as shown in Figure 1D. Use straight flat plates to attach the webcams to the T-slotted profile.
  13. Mount the UV lamps on the inside, the front, and the back profiles, and attach to the straight flat plates as shown in Figure 1D.
  14. Connect the four webcams to the USB ports of a computer running a video surveillance software.

2. Animal housing prior to experimentation

  1. House experimental mice, either purpose-bred or obtained from an external site, in groups of four. When possible, use age-matched female or male mice for these experiments. If animals are obtained from an external source, allow them to acclimatize for at least 7 days before conducting any experimental procedures.
  2. Throughout their time in the animal facility, house the animals in standard cages containing bedding and enrichment (e.g., plastic igloo, wheel, piece of paper for shredding) and keep them under a 12 h day/night cycle, with access to water and dry mouse chow ad libitum.

3. RT-VSA recordings during the light and dark phases of the day

NOTE: The protocol below describes the use of RT-VSA to assess mouse voiding behavior during the light and dark phases of the day. The animals are held on a 12 h light and 12 h dark cycle with Zeitgeber time (ZT) = 0 at 07:00 a.m. Recordings start between 10:30 a.m. and 11:00 a.m. (ZT = 3.5–4.0) for the light phase experiments and between 06:00 p.m. and 06:30 p.m. (ZT = 11.0–11.5) for the dark phase experiments. When animals are tested under both conditions, experiments are typically performed on two separate days, with at least 5 consecutive days between the light and dark tests. Experiments should not be performed on days when the animal rooms are cleaned or the cages are changed, as these can result in stresses that affect voiding behavior. All steps should be performed under conditions of minimal stress for the mice.

  1. Transport the experimental animals from their housing location to the procedure room where the RT-VSA recording chambers are located.
  2. RT-VSA recording chamber preparation
    1. Place a piece of filter paper (24.3 cm x 36.3 cm) in the bottom of each RT-VSA recording cage. Depending on the time of the day, use thin (light phase experiments) or thick (dark phase experiments) filter paper.
      NOTE: Thick filter paper is used during the active dark phase as it is more resistant to shredding than thin filter paper. Top cameras are used to visualize urine spots deposited on thick filter paper during the dark phase. In contrast, during the light phase, the bottom cameras are used because the ambient light prevents the top cameras from detecting urine spots deposited on the filter paper. In this case, thin filter paper is used. We found that the bottom cameras are ineffective at detecting small voiding events deposited on thick filter paper.
    2. On top of the filter paper, place the following items: a plastic igloo (which affords a sleeping space), a sterile 1.5 mL micro-centrifuge tube for enrichment purposes, and a 60 mm x 15 mm plastic dish containing two or three pieces of mouse dry chow and 14–16 g of water in the form of a gel-pack (non-wetting water gel; Figure 2).
      NOTE: The use of 1.5 mL micro-centrifuge tubes for enrichment purposes was specific to the enclosure used in this study.
    3. Once the recording chambers are ready, gently place the experimental mice inside by laying them down softly on the filter paper. Make sure the transfer of the animals from the housing cage to the recording cages occurs with minimum stress.
    4. Once all the experimental animals are inside their recording cages, with their lids closed, cover the top of the lids with an absorbent bench bluepad to minimize direct ambient light reflections on the plexiglass lid surface.
    5. Turn on the UV lights in the lower chamber.
      NOTE: The animals have no direct contact with the UV light. The recommended UV light is detailed in the materials section.
  3. RT-VSA recordings
    1. To record video from the top and bottom cameras, use a video surveillance recording software, which can be configured to record from multiple webcams or networked cameras at one time.
    2. Upon opening the program, initiate recording by pressing Command and R in the program window. Perform video recordings at a rate of 1 frame per s.
    3. Immediately after initiating the recordings, exit the room, closing the door gently. Ensure that the room remains quiet for the entire duration of the experiment.
  4. Finish RT-VSA recordings
    1. Return to the procedure room after 7 h (light phase experiments) or the next morning (dark phase experiments).
    2. Stop the recordings by pressing Command and T. Turn off the UV lights.
    3. After stopping the recording, the software automatically generates a movie file (in .m4v format) for each camera and saves it under the camera’s name in a previously selected destination folder. Verify that, within each camera folder, the experiments are organized into folders by date.
    4. In each date/experiment folder, verify that there is one .m4v file and all the individual .jpeg files that correspond to each of the movie frames.
    5. NOTE: The .jpeg files can be used to recover the experiment in case movie files get corrupted.
    6. Create a folder on the desktop with the name and date of the experiment and transfer all the .m4v files into this folder. If required, delete the .jpeg files once the movie files are saved and backed up.
    7. NOTE: For light phase experiments, the software will generate one movie per camera, while for dark phase experiments, it will generate two movies for each camera. This is because a new .m4v file is generated after midnight when the date changes.
    8. Copy the folder containing the movies into a flash drive for analysis in an external computer. This step can take several minutes and can be performed in parallel to steps 3.5.1 to 3.5.3.
  5. Cleaning of recording cages and transfer of experimental mice back to their housing location
    1. Remove the bluepad covering the lids of the cages. Transfer the animals from the recording cages to their housing cage.
    2. Remove the accessories and foodstuffs in the recording chambers (i.e., plastic igloos, plastic tubes, dish with rests of chow and water gel, and filter paper) and dispose of them in biohazardous waste.
    3. Clean the cages using a hand vacuum cleaner, removing chow and fecal pellets present on the bottom of the cage. Then, spray the floor and internal walls of the cage with 70% ethanol and clean the interior with a piece of soft cloth. Leave the lid of the cages open to allow them to air dry.
    4. Place the housing cage with the animals in a secondary container and transport the animals back to their housing location.

4. Generation of calibration curves

NOTE: A calibration curve is needed to convert void spot areas into urine volumes. If performing experiments during the light and dark phases of the day, then two calibration curves should be generated, one for each type of filter paper used (thin and thick filter papers). Calibration curves are generated in duplicate. Each replicate is run on a filter paper placed in a RT-VSA recording chamber. Given its complex composition, and UV excitability, use mouse urine to make the calibration curves.

  1. Mouse urine collection
    1. Take a piece of flexible transparent film (10 cm x 15 cm) and place it on a bench.
    2. Pick a mouse by its tail and scruff it. Softly massage the lower abdomen to induce urination. Collect urine on the surface of the transparent film plastic sheet.
    3. Release the animal gently inside its cage. Using a pipette, transfer the urine from the surface of the transparent film to a sterile 1.5 mL micro-centrifuge tube. Repeat the procedure with multiple mice until ~10 mL of mouse urine is collected, pool the urine, and store at -20 °C.
      NOTE: A total of ~10 mL of mouse urine is required to generate duplicate calibration curves for the light and dark phases. To avoid stressing experimental mice, do not use mice that will be subjected to RT-VSA experiments for urine collection.
  2. Calibration curve recordings
    1. Thaw the urine collected in step 4.1 and mix it by gently vortexing for 10–15 s.
    2. Place a piece of thin filter paper (24.3 cm x 36.3 cm) in each of the two RT-VSA recording cages.
    3. Pipette the following urine volumes (in μL) on each of the filter papers: 2, 5, 10, 25, 50, 80, 100, 200, 300, 400, 500, and 750. To prevent spot overlap as a result of diffusion, allocate the spots at a sufficient distance from each other.
    4. Close the lid of the cages and cover them with pads.
    5. Start recording by pressing Command and R, applying the same software parameters used to record the experiments (step 3.3.).
    6. Register the calibration curve for 1 h to allow maximal diffusion of the urine spots. Press Command and T to stop recording.
    7. Create a new folder in the desktop, place the .m4v files in it, and then transfer the data to a flash drive for subsequent analysis.
    8. Perform a similar procedure to generate duplicate curves with thick filter paper. In this case, add extra layers of pads to darken the interior and simulate dark phase conditions.
  3. Analysis of the calibration curve recordings
    1. Open the .m4v files in a movie player software, maximizing the window to fill the screen. To analyze the calibration curves performed on thick filter paper, use the files obtained with the top cameras (Figure 3A). To analyze the calibration curves performed on thin filter paper, use the files obtained with the bottom cameras (Figure 3B).
    2. Play the .m4v file, moving the time slider forward and backward to get an overview of the complete 1 h calibration curve movie.
    3. Identify the time range where the smaller urine spots (<25 μL) have the greatest intensity and have spread maximally. Take a screenshot within this time range. Name the screenshot file and save it as a .png file (Figure 3A, upper panel).
      NOTE: Screenshots are taken by pressing the key F6. To set up F6 for screenshot acquisition, select: System Preferences > Keyboard > Shortcuts, and write F6 in the box found to the right of Save Picture of Screen as a File option.
    4. Identify the time range where the medium and large urine spots (>50 μL) have maximal area and take a screenshot. It is possible that some of the smaller urine spots will not be visible by the time the larger urine spots exhibit maximal diffusion. Name the file and save the screenshot as a .png file (Figure 3A, lower panel).
    5. Open the ImageJ software (NIH) and then drag and drop the .png file icon obtained in step 4.3.3 to open the image file (Figure 4A).
    6. From the toolbar, select the Polygon Selections icon, and delineate the border of the filter paper.
    7. Then, select Analyze from the menu bar, choose Set Measurements from the expanded menu, and from the window that pops up select Area. Click OK. This allows to obtain values of area when step 4.3.8 is performed.
    8. Next, select Analyze again from the menu bar and choose Measure from the expanded options. A results window pops up containing a column area which shows the area values in pixel2 (Figure 4B–D).
    9. Select the Freehand Selections icon from the toolbar and use it to draw a line around the perimeter of an individual void spot. Measure the area as in step 4.3.8. The software updates the results table as new measurements are performed. The new set of numbers appears below the previous ones. Record the number that appears under the column area of the results window for each spot analyzed (Figure 4E–G).
    10. Repeat steps 4.3.5 to 4.3.7 for each of the replicate spots.
    11. Set the total area of the filter paper as 100% and calculate the percentage of area (% area) for each urine spot. This normalization will correct errors that might occur as a consequence of differences in the zoom or positioning of the cameras.
    12. Create a new XY table in a graphing program and insert the values of urine volume (in μL) in the X column and the duplicate values of % area in the Y column.
    13. Then, select Analysis > Analyze > XY analysis > Nonlinear Regression (curve fit). From the parameters window that appears, select Model > Polynomial > Second Order Polynomial (quadratic).
    14. Then, from the method tab, click to mark the following selections: Least Squares Regression, No Weighting, and Consider Each Replicate Y Value as an Individual Point.
    15. From the constrain tab, select for B0, Constraint Type > Constant equal to, and type 0 under the Value column. Click OK.

5. Analysis of the experimental mice recordings

  1. Open a movie file collected during the light phase (bottom camera) or dark phase (upper camera) for analysis.
  2. Assess the quality of the movie file by moving the time scroller forward and backward, confirming that the filter paper remains intact (with no tearing or chewing) during the 6 h time window to be analyzed. If the paper is torn, perform no further analysis, as the mouse might have urinated on the exposed plastic which cannot be quantified (Figure 5).
  3. To analyze experiments collected in the light or dark phases, use the fast-forward command or the time bar slider to move to the desired time window. Voiding activity during the light phase is recorded between 11:00 a.m. and 05:00 p.m. (ZT = 4.0–10.0) and during the dark phase between midnight to 06:00 a.m. (ZT = 17.0–23.0).
  4. Play the movie in fast-forward mode by clicking on the >> icon (or manually scroll through the movie), looking for evidence that the mouse is voiding. The easiest way to tell that this is occurring is to look for the sudden appearance of bright spots of urine on the filter paper. Another indicator is to look for behavioral changes including movement to the corners of the cage and a brief period of inactivity when the mouse is voiding.
    NOTE: As one becomes better at detecting voids, one can increase the speed of scrolling or fast forwarding. However, in mice with bacterial and chemically induced cystitis, which have very large numbers of small voids4,34, one can miss voiding events if one moves too quickly through the movie.
  5. Register the time at which each void occurs. As a convention, the time of the void is recorded at the first sign that urine is detected (Figure 6A,B).
  6. To make measurements of the void, first use the scroll bar to move forward (or backward) in time, looking for the point in time when maximal diffusion of the urine spot has occurred. Pause the movie at this point, and take a screenshot as described in step 4.3.3. Place the computer mouse arrow at the spot under analysis, so the spot of interest is marked in the screenshot (Figure 6C,D).
  7. Name the screenshot file using correlative numbers to account for the order of appearance in the movie.
  8. Continue analyzing the file, repeating steps 5.5 and 5.7 for each void spot on the movie. Once all the void spots have been analyzed, measure the total area of the filter paper by capturing a screenshot and using the steps described in 4.3.6.
  9. Calculate the % area of each of the urine spots as described in step 4.3.9. Transform the values of % area into volume of urine (μL) for each void spot using the calibration curves generated in step 4 and the interpolate function in the graphing software.
    1. In the graphing software, open the XY table containing the data of the calibration curve and insert the % area values in the Y column below the last value of the calibration curve (Figure 7A).
    2. Click on the Table of Results tab, and under the model tab, click to select Interpolate Unknowns from Standard Curve and press OK. A tab named interpolated X mean values appears next to the table of results tab. This tab contains a table with the interpolated values that correspond to the volume of urine in μL for each void spot. (Figure 7A,B).
  10. Repeat steps 5.1 to 5.9 to analyze all the experimental mice.
  11. Create a workbook file that contains the data obtained from steps 5.5 to 5.10, using one spreadsheet per mouse (Figure 7C). Create one file for the light phase experiments and another for the dark phase ones. These master files contain all the raw data and necessary calculations used in further analyses.

6. Analysis of the urination pattern of experimental mice

  1. Generate primary and secondary void spot profiles (Figure 8A,B and Figure 9).
    NOTE: According to frequency distribution studies23, void spots that are ≥20 μL typically represent >95% of the total voided volume and are considered primary void spots (PVSs)8,23,35. Void spots that are ≤20 μL are considered secondary or small void spots (SVSs). Discrimination between PVSs and SVSs has been shown to be a useful approach to characterize voiding phenotypes. An elevated number of SVSs indicates voiding dysfunction35.
    1. From the master file containing voiding spot data of interest (light phase or dark phase), classify the spots based on their volume as PVSs when their volume is ≥20 μL, or SVSs when their volume is <20 μL.
    2. Count the number and calculate the average voided volume and the total volume for the PVSs. Count the number and calculate the total volume for the SVSs.
    3. Generate a graph bar that compares the control to the treated mice for each of the calculated parameters: number of PVSs, voided volume of PVSs, total volume of PVSs, number of SVSs, total volume of SVSs.
  2. Optional: Generate a cumulative voided volume plot (staircase function) to show voiding behavior over time (Figure 10 and Figure 11).
    1. Open the workbook master file and convert the time of voiding, which is expressed in hours, minutes, and seconds, into the decimal form. Calculate the cumulative urine volume (in μL) for each time point.
    2. Generate an XY table in the graphing software with time in decimal form in the X column and cumulative urine volumes in the Y column. Copy the time and urine volume data into the table. Add (0; 0) and (6; maximum value) data points. These points are necessary to complete the horizontal lines of the plot at the start of the experiment (time: 0 h) when the voided volume is zero (0; 0), and at the end of the experiment (time: 6 h) when the cumulative value of the voided urine is equal to the value obtained for the last voiding event (6; maximum value).
    3. Double click on the graph; the format graph window should appear. Select the Appearance tab, click to unselect the button for Show Symbols and click to select Show Connecting Line/Curve. Click on the Style option and select Survival.

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Results

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Voiding behavior of urothelial Piezo1/2 knockout mice

During the storage phase of the micturition cycle, the urothelium is hypothesized to sense the tension exerted by the urine accumulated in the bladder and to transduce this mechanical stimulus into cellular responses such as serosal ATP release1,3. We have previously shown that mechanically activated PIEZO1 and PIEZO2 channels are expressed in the mouse uro...

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Discussion

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The incorporation of video-monitoring is a cost-effective modification that presents several advantages over the classical VSA. In the classical VSA, which is typically used as an end-point assay, it is difficult to distinguish overlapping void spots. This is not a trivial concern, as mice tend to urinate multiple times in the same area when the assay is prolonged for several hours, typically in the corners of their cage. Thus, the first advantage of RT-VSA is that the investigator can readily identify individual spots t...

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Disclosures

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The authors have nothing to disclose.

Acknowledgements

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This work was supported by an NIH grant R01DK119183 (to G.A. and M.D.C.), a pilot project award through P30DK079307 (to M.G.D.), an American Urology Association Career Development award and a Winters Foundation grant (to N.M.), and by the Cell Physiology and Model Organisms Kidney Imaging Cores of the Pittsburgh Center for Kidney Research (P30DK079307).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1.00” X 1.00” T-Slotted Profile - Four Open T-Slots –  cut to 10 inches80/201010Amount: 20
1.00” X 1.00” T-Slotted Profile - Four Open T-Slots –  cut to 12 inches80/201010Amount: 6
1.00” X 1.00” T-Slotted Profile - Four Open T-Slots –  cut to 40 inches80/201010Amount: 4
1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 14.75 inches80/201010Amount: 12
1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 32 inches80/201010Amount: 5
1/4-20 Double Slide-in Economy T-Nut80/203280Amount: 16
1/4-20 Triple Slide-in Economy T-Nut80/203287Amount: 18
10 & 25 Series 2 Hole - 18mm Slotted Inside Corner Bracket with Dual Support80/2014061Amount: 6
10 Series 3 Hole - Straight Flat Plate80/204118Amount: 8
10 Series 5 Hole - "L" Flat Plate80/204081Amount: 8
10 Series 5 Hole - "T" Flat Plate80/204080Amount: 8
10 Series 5 Hole - Tee Flat Plate80/204140Amount: 2
10 Series Standard Lift-Off Hinge - Right Hand Assembly80/202064Amount: 2
10 to 15 Series 2 Hole - Lite Transition Inside Corner Bracket80/204509Amount: 6
24”-long UV tube lightsADJ Products LLCT8-F20BLB24Amount: 2
20W bulb – 24” Wavelength: 365nm
Acrylic Mirror SheetProfesional PlasticsAmount: 1
82.5 cm x 26.5 cm
Acrylic Mirror SheetProfesional PlasticsAmount: 2
26.5 cm X 30.5 cm
Acrylic Mirror SheetProfesional PlasticsAmount: 2
82.5 cm x 30.5 cm
AR polycarbonate (UV resistance)80/2065-2641Amount: 2
4.5mm Thick, Clear, 38.5 cm x 26.5 cm
AR polycarbonate (UV resistance)80/2065-2641Amount: 4
4.5mm Thick, Clear, 38.5 cm x 21.5 cm
AR polycarbonate (UV resistance)80/2065-2641Amount: 4
4.5mm Thick, Clear, 26.5 cm x 21.5 cm
AR polycarbonate (UV resistance)80/2065-2641Amount: 4
4.5mm Thick, Clear 37.5 cm x 23.9 cm
AR polycarbonate (UV resistance)80/2065-2641Amount: 4
4.5mm Thick, Clear , 24.4 cm x 23.9 cm
Chromatography paper (thin paper) Thermo Fisher Scientific57144
Cosmos blotting paper (thick paper)Blick Art Materials10422-1005
ExcelMicrosoft Corporation
GraphPad PrismGraphPad SoftwareVersion 9.4.0graphing and statistics software
ImageJ FIJINIH
ParafilmMercktransparent film
Quick Time Player 10.5 software Applemultimedia player
Security spyBen softwarevideo surveillance software system
Standard End Fastener, 1/4-2080/203381Amount: 80
UV transmitting acrylicSpartechPolycast Solacryl SUVTAmount: 2
38.5 cm x 26.5 cm 
Water gel: HydroGelClearH2O  70-01-5022(https://www.clearh2o.com/product/hydrogel/)
WebcamLogitechC930eAmount: 4

References

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Void Spot AssayReal Time VSAVoiding BehaviorMouse ModelsUrine Spot AnalysisVoluntary MicturitionVideo MonitoringBladder FunctionUrethral SphincterCyclophosphamide Treatment

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