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.
Urine storage and micturition are coordinated by a central circuitry 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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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)
- 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).
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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).
- 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.
- 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.
- 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.
- 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.
- 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.
- Connect the four webcams to the USB ports of a computer running a video surveillance software.
2. Animal housing prior to experimentation
- 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.
- 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.
- Transport the experimental animals from their housing location to the procedure room where the RT-VSA recording chambers are located.
- RT-VSA recording chamber preparation
- 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.
- 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).
- 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.
- 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.
- Turn on the UV lights in the lower chamber.
- 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.
- RT-VSA recordings
- 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.
- 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.
- 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.
- Finish RT-VSA recordings
- Return to the procedure room after 7 h (light phase experiments) or the next morning (dark phase experiments).
- Stop the recordings by pressing Command and T. Turn off the UV lights.
- 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.
- 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.
- NOTE: The .jpeg files can be used to recover the experiment in case movie files get corrupted.
- 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.
- 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.
- 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.
- Cleaning of recording cages and transfer of experimental mice back to their housing location
- Remove the bluepad covering the lids of the cages. Transfer the animals from the recording cages to their housing cage.
- 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.
- 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.
- 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.
- Mouse urine collection
- Take a piece of flexible transparent film (10 cm x 15 cm) and place it on a bench.
- 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.
- 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.
- Calibration curve recordings
- Thaw the urine collected in step 4.1 and mix it by gently vortexing for 10–15 s.
- Place a piece of thin filter paper (24.3 cm x 36.3 cm) in each of the two RT-VSA recording cages.
- 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.
- Close the lid of the cages and cover them with pads.
- Start recording by pressing Command and R, applying the same software parameters used to record the experiments (step 3.3.).
- Register the calibration curve for 1 h to allow maximal diffusion of the urine spots. Press Command and T to stop recording.
- 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.
- 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.
- Analysis of the calibration curve recordings
- 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).
- Play the .m4v file, moving the time slider forward and backward to get an overview of the complete 1 h calibration curve movie.
- 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.
- 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).
- 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).
- From the toolbar, select the Polygon Selections icon, and delineate the border of the filter paper.
- 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.
- 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).
- 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).
- Repeat steps 4.3.5 to 4.3.7 for each of the replicate spots.
- 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.
- 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.
- Then, select Analysis > Analyze > XY analysis > Nonlinear Regression (curve fit). From the parameters window that appears, select Model > Polynomial > Second Order Polynomial (quadratic).
- 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.
- 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
- Open a movie file collected during the light phase (bottom camera) or dark phase (upper camera) for analysis.
- 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).
- 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).
- 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.
- 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).
- 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).
- Name the screenshot file using correlative numbers to account for the order of appearance in the movie.
- 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.
- 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.
- 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).
- 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).
- Repeat steps 5.1 to 5.9 to analyze all the experimental mice.
- 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
- 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.
- 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.
- 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.
- 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.
- Optional: Generate a cumulative voided volume plot (staircase function) to show voiding behavior over time (Figure 10 and Figure 11).
- 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.
- 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).
- 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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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 urothelium3,36. To determine if urothelial PIEZO channels are important for normal voiding behavior, we performed RT-VSA on urothelial-conditional Piezo1/2 double knockout mice (Pz1/2-KO) mice and age- and sex-matched controls (Pz1/2-C; Figure 9). We tested both female and male mice for 6 h during the dark and light phases of the day, and analyzed the data as described in step 6.1. When we compared the voiding activity between the light and dark phases of the day for the control groups, it was noticed that both females and males exhibited an increase in the number and total volume of PVSs during their active period. This observation indicates that, even though the mice were in a foreign environment, the circadian nature of mouse voiding behavior (highest during their active dark phase) was preserved. During their inactive light phase, we did not observe any significant differences in any of the parameters analyzed for female or male Pz1/2-KO mice when compared to the control counterparts (Pz1/2-C). However, when we tested the animals during the active dark phase, both female and male Pz1/2-KO mice showed an altered voiding phenotype, characterized by a significant increase in the number and in the total volume of SVSs (Figure 9). There were no significant differences in PVS number, average volume per PVS, or total PVS volume in female or male Pz1/2-KO compared to their corresponding controls. These results indicate that urothelial PIEZO1/2 channels do not play a significant role in voiding function during the mouse inactive light phase but are important in preventing frequent spotting during their active dark phase.
The results presented in Figure 9 illustrate two of the advantages of employing RT-VSA versus the endpoint VSA. First, if the analysis was performed only during the light phase, we would have misconcluded that urothelial PIEZO1/2 channels had no role in voiding function. Second, in most of the endpoint VSA studies, SVSs were treated as experimental noise, and in some cases excluded from subsequent analysis23. In the RT-VSA, we could determine that the SVSs were individual events, not secondary to PVSs (carryover or end micturition dribbling) and were voluntary in the sense that the animals moved to the periphery of the cage, voided, and then left. This observation provided us with valuable information about the phenotype of Pz1/2-KO mice that otherwise would have been missed.
Voiding behavior of female mice under basal conditions and following cyclophosphamide treatment
In this pilot experiment, we tested the effects of cyclophosphamide (CYP) on mouse voluntary voiding behavior. Cyclophosphamide, a drug used to treat some forms of cancer or immunological disorders, is known to cause cystitis in patients and experimental animals, resulting in an overactive bladder phenotype characterized by multiple, small voiding events26,37,38. To determine the baseline voiding behavior, we placed an untreated female mouse in the RT-VSA during the dark phase. The same mouse was injected intraperitoneally with a dose of CYP (150 mg/kg) 5 days later to cause acute cystitis. Immediately after injection, the mouse was placed in an RT-VSA cage for recording. The voiding behavior is represented in a step plot (Figure 11), with the vertical lines representing the volume of urine voided and the horizonal lines representing the time. Compared to the results obtained under basal conditions, it is evident that the amount of urine released per void is smaller after CYP treatment, and that the micturition events are much more frequent. This can be better appreciated in the inset of the graph, where the first 30 min of the analyzed period (corresponding to midnight to 00:30 a.m. [7.0 to 7.5 h post injection]) is magnified.
Figure 1: The RT-VSA recording chamber. (A) The RT-VSA recording system includes an upper portion, which is comprised of two side-by-side, clear, acrylic chambers and associated upper cameras. The lower half of the device is a single unit stand that holds the upper mouse chambers and contains the bottom cameras and UV lights. The interior of the walls of the stand are made of mirror panels that reflect the UV light to provide even illumination to the bottom of the cages. Not depicted is a computer that receives a video feed from the cameras and records the data. (B) Model of the mouse chamber with components and dimensions. 1) standard lift-off hinge; 2) camera mounting bracket; and 3) lite transition corner bracket. Front (C) and top (D) views of the UV stand with components and dimensions. UV lights are mounted in the interior of the front and back profiles with straight flat plates. A 32 in profile secured to the stand main frame with two lite transition corner brackets is used to mount the cameras. Straight flat plates are used to attach the cameras to the 32 in profile. A mirrored panel is mounted in the interior of the stand (not shown). 1) five-hole tee flat plate; 2) five-hole T flat plate; and 3) five-hole L flat plates. All values presented are in inches. Please click here to view a larger version of this figure.
Figure 2: RT-VSA mouse chamber. Prior to experimental procedures, each experimental mouse chamber is prepared by lining it with a piece of filter paper (numbered 1). Depending on the period of the day when the experiment is performed, the filter paper used is either thick (dark phase) or thin (light phase). A plastic dome is placed in the center of the chamber (numbered 2), a plastic dish with water (in the form of non-wetting water gel) and chow is placed to one side of the dome (numbered 3), and a plastic 1.5 mL micro-centrifuge tube, a form of enrichment, is placed on the opposite side (numbered 4). The initial arrangement of elements within the cage is kept consistent in all experiments. Notice that a piece of filter paper is placed between the two neighboring cages and is marked in the figure with an asterisk. This is to prevent the influence of visual cues arising from the neighboring mouse. Please click here to view a larger version of this figure.
Figure 3: Calibration curve. Representative images of thick (A, left panels) and thin (B, left panel) filter papers spotted with known amounts of mouse urine. Images are screenshots from video recordings obtained using the top (A, left panels) or bottom (B, left panel) cameras. Red arrows point to the 2 µL urine spots, and the numbers below the spots are the spot volume values in µL. Notice that the small urine spots (<50 µL) in the upper image of panel A that are visible shortly after urine spotting (5 min) are no longer noticeable after 30 min (lower panel). Right panels: Plot of void spot area (expressed as the percentage of total filter paper area) as a function of the urine volume. As noted by others, the data of void spot area as a function of the urine volume does not follow a linear relationship30. Thus, we fit the data to a second order polynomial using nonlinear regression. Data were constrained such that X = 0 to Y = 0. Please click here to view a larger version of this figure.
Figure 4: Determination of filter paper and urine spot areas using ImageJ. (A) A screenshot of the calibration curve shown in Figure 3A opened in ImageJ. The boxed region in (A) is shown in more detail in (B). (B) The polygon selections icon is selected (circled in red) to draw a polygon along the perimeter of the filter paper (partial view; yellow line with red arrows pointing to the nodes generated by the selections tool). (C) The area of the polygon reflects the total area of the filter paper and is measured by selecting Analyze > Measure (red circle). (D) The value of the area (in pixels) appears in a results window (red circle). (E) To measure the area of an individual void spot, the freehand selections tool is chosen (red circle) and in (F) is used to draw a line around the border of the spot (the red arrow points to the spot). Using the same command as in (C), the area of the void spot is measured, with the result appearing as a new value in the results window, immediately below the last measurement taken (panel G, red circle). Please click here to view a larger version of this figure.
Figure 5: Assessment of filter paper status after finishing RT-VSA experiments. Representative images of the mouse chamber showing different conditions of the filter paper during or after an RT-VSA experiment. Filter paper that remained intact until the end of the testing period is shown in (A) and represents the most common situation and the condition under which an experiment can be further analyzed (green checkmark). Examples of damaged filter paper are shown in (B) and (C) ,and are representative of experiments that must not be further analyzed (red X). In (B), the filter paper was shredded by the mouse (red arrow) in one of the corners. The dashed yellow line marks the border of the cage’s floor. The boxed region is magnified in the image to the right. In (C), the integrity of the experiment was compromised as a result of non-wetting water gel (red arrow) being moved by the mouse into the corner, leaving three large water spots (red asterisks). These movements were confirmed by watching the video. Images are screenshots from movie files obtained during the dark phase, using the top camera, and with the cage lined with thick filter paper. Please click here to view a larger version of this figure.
Figure 6: Determination of the time of voiding and acquisition of screenshots for void spot area measurements. (A,B) Screenshot of a mouse in a corner of its chamber, with a void spot that has just become visible. The time of this frame is annotated as the time of voiding. (C,D) Once the void spot reaches maximal diffusion, a screenshot is taken to determine the volume of the void. Images are screenshots from movie files obtained with the top camera, using thick filter paper, and during the dark phase. The period of analysis was midnight to 06:00 a.m. Please click here to view a larger version of this figure.
Figure 7: Transformation of void spot area into urine volume and generation of a worksheet file with experimental raw data. (A) Screenshot of the graphing software window showing values for void spots expressed as the percentage of total filter paper area (data outlined in green). The worksheet (green arrow) is saved under the data tables section. (B) Conversion of the percentage of total area to urine volume (data outlined in red). The latter values are shown in the results section (red arrow) and are calculated from the calibration curve using interpolate unknowns from standard curve function. (C) Screenshot of a worksheet with compiled raw and calculated data. Please click here to view a larger version of this figure.
Figure 8: Classification of void spots into PVSs and SVSs and their analysis. Screenshots of worksheets (generated as shown in Figure 7C) from a mouse under basal conditions (A) and 7 h after receiving a dose of cyclophosphamide (150 mg/kg) (B). Void spots are classified as primary void spots (PVSs) if the volume is ≥20 µL, or as small void spots (SVSs) if the volume is <20 µL. Note that in the untreated mouse (A), all voiding events are PVSs, while in the cyclophosphamide-treated animal, most voids are SVSs. The circled table in each panel (red outline), include the following summary statistics: number of voids, average voided volume, and total voided volume. Please click here to view a larger version of this figure.
Figure 9: Voiding behavior of Piezo1/2 knockout and control mice. The voiding behavior of female (A) and male (B) Piezo1/2 knockout (Pz1/2-KO) mice and control counterparts (Pz1/2-C) was recorded during a 6 h time window during the dark or light phase of the day, and results analyzed as described in step 6.1. Data are shown as mean ± standard error of mean (S.E.M.). Data were compared using a Mann-Whitney test. Abbreviations: PVSs = primary void spots; SVSs = small void spots; NS = not significant; * p < 0.05; ** p < 0.01. This figure has been modified from 3. Please click here to view a larger version of this figure.
Figure 10: Worksheets for the calculation and graphical representation of the cumulative voided volume as a function of time. (A) Screenshot of the worksheet generated in Figure 7C displaying the calculations needed to generate a staircase function plot of the cumulative volume of urine voided as a function of time. Time must be entered in a decimal format (column G, red square). To transform the time to decimal format, three columns (red arrows) were added to the right of the time of voiding column to separately annotate the values of hours (column D), minutes (column E) and seconds (column F), and the final result of time in a decimal format (column G). To transform the time voiding to a decimal format, use the following function: (h) + (min/60) + (s/3,600). One column was added to the right of the urine volume column, to display the cumulative urine volume (blue rectangle), which is calculated as the sum of all urine volumes (column J) obtained up to the time of a new voiding event (including the new event value). (B) Screenshot showing data copied from the worksheet in panel A and pasted into column X (time in decimal format) and column Y (cumulative urine volumes) in a graphing software project. The first and last time points of the plot are set to 0 and 6 (maximum value) rows as indicated by the red arrows. (C) To generate a step plot, the graph is formatted by deselecting the show symbols field, enabling the show the connecting line/curve option, and selecting the survival style, as shown in the fields encircled in red. Please click here to view a larger version of this figure.
Figure 11: Representative results of mouse voiding behavior under basal conditions or after cyclophosphamide administration. The voluntary voiding behavior of a mouse was assessed by RT-VSA during the dark phase before (basal) and 7 h after cyclophosphamide (CYP) administration, as described in the Representative Results. The time and volume of the voiding events is represented in a step plot that was generated as described in step 6.2. The inset of the graph represents the area of the plot marked with a green rectangle and corresponds to the first 30 min of the period analyzed. Please click here to view a larger version of this figure.
Figure 12: Identification of overlapping urine spots by RT-VSA. Screenshots of video frames captured with the top camera in the dark phase of a RT-VSA experiment (images 1 and 2). The boxed region in image 2 is magnified in image 3. The mouse voided in the same upper right corner two additional times during the next few hours (images 4–6). Despite the overlap of void spots, the individual voiding events were readily detected using RT-VSA. To visualize how these events would appear as an endpoint assay, we exposed the corner of the filter paper to different lighting conditions. Under visible, ambient light, only one single, large void spot could be identified (image 7). Likewise, if this same region was examined under UV light, only a single dark spot was distinguishable (image 8). Thus, multiple voids in the same region are readily detected by RT-VSA, but such discrimination is not possible if the analysis is performed as an end-point assay. Please click here to view a larger version of this figure.
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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 that have been deposited partially or totally on top of each other. This is well illustrated in the experiment shown in Figure 12. In this case, video recording confirms that the mouse voided three times in the same corner, depositing overlapping void spots. If the filter paper was analyzed only at the end of the experiment, in the manner typical of a VSA, only one spot could be distinguished. So, while the classical, endpoint VSA can be useful in identifying large differences in voiding behavior, it may fail to discriminate between more subtle phenotypes. With the current advances in genetics and the generation of transgenic animals, the development of methods that can accurately assess voiding behavior is essential.
Second, RT-VSA provides the investigator with temporal information about the voiding events. Thus, one can determine true micturition frequencies, as well as establish relationships between voiding events. For example, in a situation where a cluster of SVSs is observed next to a single large PVS, in the classical VSA, one cannot readily discriminate if the mouse voided several times during the experiment (with three small and one large void) or if the animal deposited one large urine volume, and the three smaller spots are a result of post-micturition dribbling or carryover from fur. RT-VSA can be used to address these questions.
Third, RT-VSA allows one to analyze the behavior of a mouse before, during, and after the voiding event. The investigator can also monitor the mouse’s overall activity profile by analyzing RT-VSA recordings with freely available software such as Mouse Behavior Tracker39. In addition, one can discriminate whether an animal stops to urinate (typical behavior of females), whether the animal is in motion during voiding (which in males can result in the trailing of void spots), or whether it voids during its sleep/rest phase. We have observed the latter behavior, although it is normally rare. Thus, if properly validated and interpreted, observing voiding behavior can provide the investigator with valuable insights into voiding dysfunction, including, for example, nocturia or incontinence. RT-VSA (and VSA) allows investigators to determine the fraction of void events that occur in the central area of a cage, as opposed to the corners. Normally, center voiding is rare, and increased numbers of such events is a sign of bladder dysfunction8,22. The one exception is dominant alpha-males, which urinate frequently and across the entirety of their enclosures20. In order to perform an analysis of central voiding using RT-VSA, one should consider moving the food dish to a side wall and remove the dome (or fix it to a particular region of the cage), so spatial patterns of urination could be more readily established.
Fourth, the RT-VSA allows one to more effectively manage the stresses associated with cage exchange compared to classical VSA. Mice are skittish by nature and handling them or transferring them to a new environment causes stress, which is known to impact voiding behavior35. By its nature, it is difficult to incorporate a period of acclimatization into the standard VSA. In contrast, in RT-VSA, a period of acclimatization can be easily incorporated in each experiment, and any voiding events during this period disregarded during the analysis. An additional stress-relieving aspect of RT-VSA implemented in this protocol is the duplication of conditions under which mice are normally housed (i.e., access to food and water ad libitum, a dome, and a toy). This is, to the best of our knowledge, one of the richest environments described for this type of experiment. In a typical endpoint VSA, mice are deprived from water (and sometimes also from food)8,22, which we provide ad libitum.
Fifth, in addition to the positive effect that this enriched environment may have on the stress levels of mice, it allows the investigator to perform RT-VSA for extended periods of time, typically in 6 h time windows; however, more recently we have been performing experiments that run for 24 h (data not shown). This is ideal when assessing the effects of circadian rhythmicity or when studying mice that may exhibit an underactive bladder phenotype, and thus urinate infrequently.
For 24 h video monitoring, we recommend using thin filter paper and analyzing the experiments using the bottom cameras. This combination of type of paper and camera is the best option that allows for sensitivity of detection in the dark and light phases. As mentioned earlier, bottom cameras cannot readily detect void spots on the thick filter paper and top cameras do not reliably detect void spots during the light phase. If performing these extended analyses, we suggest placing the animals in the recording cages at 05:00 p.m., analyzing from 06:00 p.m. on day 1 and completing the experiment at 06:00 p.m. on the following day.
While RT-VSA has many positives aspects, there are some caveats worth noting. While we make every attempt to limit stress when performing these analyses, we cannot completely replicate the mouse’s habitat, as the animals are normally group-housed. They also drink water from a bottle instead of getting water from non-wetting water gel. However, our observation that mice have fewer voids during their inactive light phase versus their active dark phase indicates to us that the mice retain their normal patterns of circadian voiding behavior40. Reflecting their easily stressed nature, we found that mice exhibit disrupted voiding behavior on the days of cage cleaning. Thus, one must limit analysis to periods of time when the animals will be less impacted by routine animal husbandry. An important factor to consider is whether the UV light has any impact on voiding function. Because our findings show a distinct circadian difference in VSA parameters between light and dark phases, which are consistent with other techniques that descrive a similar phenomenon but not using UV light40, we conclude that UV light illumination does not impact voiding function. Notice that the filter paper prevents light from passing from the lower chamber to the upper chamber.
An additional caveat is that the longer one keeps the mice in the chamber, the more likely they are to chew or damage the filter paper (irrespective of the thickness of the paper). While none of the animals disrupted the paper during the light phase, almost 30% of mice damaged the paper during the dark active phase. In the analysis of the 24 h experiments, we observed that a similar proportion (one third) of the experiments could not be analyzed due to paper damage, meaning that not only the amount of time, but also the phase of the day affects the shredding behavior of the animal. Unfortunately, in our experience, we have noticed that an animal that disrupts the cage’s paper will continue to do so even upon re-testing. As noted above, mice sometimes urinate on the bare acrylic floor, and thus loss of the paper makes the analysis incomplete and of limited value.
Further, the analysis of male voiding behavior can be more difficult as they tend to void more frequently, they can dribble after voiding, and there is a population of males (20%) with a more aggressive alpha-male phenotype characterized by large numbers of small voids that are distributed across the cage, as discussed in an earlier publication3. As this alpha-male behavior can constitute a confounding factor for establishing voiding phenotypes, we exclude animals with ≥50 (light phase experiments) or ≥100 (dark phase experiments) voids from analysis.
There are situations in which the discrimination of voids based on their volume (i.e, PVS vs SVS) is not necessarily warranted. For instance, in our experience, mice with bacterial or chemical cystitis tend to void smaller volumes than controls4,34. However, there are great differences in the voided volume between individual mice; in some, most voids have a volume ≤20 μL, while in others the majority is >20 μL. Consequently, the discrimination of the voided volume does not provide any advantage for the characterization of these phenotypes.
In sum, RT-VSA is an easy to implement tool for analyzing mouse voiding behavior in freely mobile mice. Unlike tools such as cystometry, it does not require surgical implantation of a catheter or non-physiological rates of bladder filling. It allows for the determination of voiding behavior in both sexes of mice, over extended periods of time, and during the light and dark phases of the day cycle. It is also relatively affordable, especially when compared to more specialized and expensive devices such as metabolic cages. While there are caveats associated with this tool, they are generally easy to manage. Finally, this technique could be easily adapted to other animal species, including other rodents.
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The authors have nothing to disclose.
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).
|1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 10 inches||80/20||1010||Amount: 20|
|1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 12 inches||80/20||1010||Amount: 6|
|1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 40 inches||80/20||1010||Amount: 4|
|1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 14.75 inches||80/20||1010||Amount: 12|
|1.00” X 1.00” T-Slotted Profile - Four Open T-Slots – cut to 32 inches||80/20||1010||Amount: 5|
|1/4-20 Double Slide-in Economy T-Nut||80/20||3280||Amount: 16|
|1/4-20 Triple Slide-in Economy T-Nut||80/20||3287||Amount: 18|
|10 & 25 Series 2 Hole - 18mm Slotted Inside Corner Bracket with Dual Support||80/20||14061||Amount: 6|
|10 Series 3 Hole - Straight Flat Plate||80/20||4118||Amount: 8|
|10 Series 5 Hole - "L" Flat Plate||80/20||4081||Amount: 8|
|10 Series 5 Hole - "T" Flat Plate||80/20||4080||Amount: 8|
|10 Series 5 Hole - Tee Flat Plate||80/20||4140||Amount: 2|
|10 Series Standard Lift-Off Hinge - Right Hand Assembly||80/20||2064||Amount: 2|
|10 to 15 Series 2 Hole - Lite Transition Inside Corner Bracket||80/20||4509||Amount: 6|
|24”-long UV tube lights||ADJ Products LLC||T8-F20BLB24||Amount: 2
20W bulb – 24” Wavelength: 365nm
|Acrylic Mirror Sheet||Profesional Plastics||Amount: 1
82.5 cm x 26.5 cm
|Acrylic Mirror Sheet||Profesional Plastics||Amount: 2
26.5 cm X 30.5 cm
|Acrylic Mirror Sheet||Profesional Plastics||Amount: 2
82.5 cm x 30.5 cm
|AR polycarbonate (UV resistance)||80/20||65-2641||Amount: 2
4.5mm Thick, Clear, 38.5 cm x 26.5 cm
|AR polycarbonate (UV resistance)||80/20||65-2641||Amount: 4
4.5mm Thick, Clear, 38.5 cm x 21.5 cm
|AR polycarbonate (UV resistance)||80/20||65-2641||Amount: 4
4.5mm Thick, Clear, 26.5 cm x 21.5 cm
|AR polycarbonate (UV resistance)||80/20||65-2641||Amount: 4
4.5mm Thick, Clear 37.5 cm x 23.9 cm
|AR polycarbonate (UV resistance)||80/20||65-2641||Amount: 4
4.5mm Thick, Clear , 24.4 cm x 23.9 cm
|Chromatography paper (thin paper)||Thermo Fisher Scientific||57144|
|Cosmos blotting paper (thick paper)||Blick Art Materials||10422-1005|
|GraphPad Prism||GraphPad Software||Version 9.4.0||graphing and statistics software|
|Quick Time Player 10.5 software||Apple||multimedia player|
|Security spy||Ben software||video surveillance software system|
|Standard End Fastener, 1/4-20||80/20||3381||Amount: 80|
|UV transmitting acrylic||Spartech||Polycast Solacryl SUVT||Amount: 2
38.5 cm x 26.5 cm
|Water gel: HydroGel||ClearH2O||70-01-5022||(https://www.clearh2o.com/product/hydrogel/)|
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