We describe the use of high frequency ultrasound with contrast imaging as a method to measure bladder volume, bladder wall thickness, urine velocity, void volume, void duration, and urethral diameter. This strategy can be used to assess voiding dysfunction and treatment efficacy in various mouse models of lower urinary tract dysfunction (LUTD).
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Liu, T. T., Rodgers, A. C., Nicholson, T. M., Macoska, J. A., Marker, P. C., Vezina, C. M., Bjorling, D. E., Roldan-Alzate, A., Hernando, D., Lloyd, G. L., Hacker, T. A., Ricke, W. A. Ultrasonography of the Adult Male Urinary Tract for Urinary Functional Testing. J. Vis. Exp. (150), e59802, doi:10.3791/59802 (2019).
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The incidence of clinical benign prostatic hyperplasia (BPH) and lower urinary tract symptoms (LUTS) is increasing due to the aging population, resulting in a significant economic and quality of life burden. Transgenic and other mouse models have been developed to recreate various aspects of this multifactorial disease; however, methods to accurately quantitate urinary dysfunction and the effectiveness of new therapeutic options are lacking. Here, we describe a method that can be used to measure bladder volume and detrusor wall thickness, urinary velocity, void volume and void duration, and urethral diameter. This would allow for the evaluation of disease progression and treatment efficacy over time. Mice were anesthetized with isoflurane, and the bladder was visualized by ultrasound. For non-contrast imaging, a 3D image was taken of the bladder to calculate volume and evaluate shape; the bladder wall thickness was measured from this image. For contrast-enhanced imaging, a catheter was placed through the dome of the bladder using a 27-gauge needle connected to a syringe by PE50 tubing. A bolus of 0.5 mL of contrast was infused into the bladder until a urination event occurred. Urethral diameter was determined at the point of the Doppler velocity sample window during the first voiding event. Velocity was measured for each subsequent event yielding a flow rate. In conclusion, high frequency ultrasound proved to be an effective method for assessing bladder and urethral measurements during urinary function in mice. This technique may be useful in the assessment of novel therapies for BPH/LUTS in an experimental setting.
Benign prostatic hyperplasia (BPH) is a disease that develops in men as they age and affects nearly 90% of men over 80 years of age1,2. Although the development of BPH is generally associated with aging, other factors including obesity and metabolic syndrome can lead to BPH in relatively younger men3,4. Many men with BPH develop lower urinary tract symptoms (LUTS) that significantly decrease their quality of life, and some experience complications that may include bleeding, infection, bladder outlet obstruction (BOO), bladder stones, and renal failure. The cost of treatment for BPH exceeds $4 billion annually5,6,7. Diagnosis of LUTS caused by BPH generally relies on the use of the AUA symptom index (AUASI) score, uroflowmetry, and assessment of prostate size8. The etiology of BPH/LUTS is complex and multifactorial, and disease development and progression has been associated with prostatic hyperplasia (prostate proliferation), smooth muscle contractility, and fibrosis. Current treatments include the use of α-adrenergic blockers to regulate smooth muscle tone within the bladder and prostate to alleviate LUTS and/or 5α-reductase inhibitors to decrease androgen metabolism and decrease prostate size. Better disease models, murine and other, to allow the accurate study of the effects of varied causative and therapeutic factors in this disease process over time is highly desirable9.
Rodent models have been extensively used to study urodynamics; however, most studies are focused on female micturition and disease10. In order to fully examine all aspects of male LUTS, rodent models have been developed and used to study different aspects of BPH including changes in cellular proliferation, smooth muscle function, collagen deposition, and inflammation11,12,13,14. However, rodent and human prostate anatomy differ. While the human prostate is compact and encased by a condensed fibromuscular layer, the rodent prostate is lobular; and these differences complicate direct comparisons of disease progression and treatment efficacy. Additionally, LUTS are difficult to assess in mice, since it is not possible to directly measure bother. Instead, current methods for studying disease correlate histological features with physiological features (i.e., bladder volume and wall thickness with uroflowmetry, void spot assays, and cystometry endpoint data) that compare the level of urinary dysfunction between BPH model and control animals12,15,16,17,18. Physiological features are frequently evaluated as post-mortem necropsy endpoints, and there is an inability within the same animal to observe BOO across time. Recently, we have identified a subdivision of the pelvic urethra (the prostatic urethra) where exogenous hormone implants cause a narrowing based on post-mortem necropsy assessments12. Current methods do not allow for the direct, in vivo assessment of urethral narrowing during voiding.
Ultrasound is a non-invasive diagnostic and evaluation technique that has successfully been used in other disease models. It is used to quantify organ volume and assess vascular flow19,20,21. Ultrasound is also used to visualize and guide microinjections, allowing for targeted injections of stem cells or other drugs, and to evaluate systolic and diastolic cardiac function.
This protocol describes the use of high frequency ultrasound to evaluate lower urinary tract anatomy and assess urinary physiology in anesthetized mice. We describe the use of ultrasound for measuring bladder volume and wall thickness. We also describe the use of contrast-enhanced ultrasound to measure urine velocity, urine volume, void duration, and urethra diameter. The use of ultrasound provides a more comprehensive understanding of the lower urinary tract in vivo, determines how disease alters normal voiding function, and gives us the tools to better evaluate the effectiveness of new therapeutic options. Currently, the non-contrast imaging protocol is non-terminal, while the current contrast-enhanced imaging protocol is a terminal procedure.
Procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Wisconsin – Madison.
1. Animal preparation
- Place a 24-month-old, C57Bl6/J male mouse in a pre-charged chamber with 3-5% isoflurane until the righting reflex is lost and the breathing rate slows.
- If necessary, use clippers to shave the abdominal hair from the animal for surgery and/or imaging. Remove all remaining hair using a depilatory cream.
- Place the mouse in a supine position in a nose cone with 2% isoflurane on a heated platform to maintain anesthesia. Confirm the depth of anesthesia by loss of motion from the animal in response to a pedal-withdrawal reflex (Figure 1B).
2. Ultrasound set-up
- Connect a MV707 probe with center frequency of 30 MHz to the active-port, with the application preset to "abdominal imaging" (Figure 1A).
- Position the ultrasound probe parallel to the long axis of the bladder (Figure 1C).
- Long and short axis images of the bladder, the prostate, and the urethra are made in B-Mode (Figure 1D).
- Use the "xy" micro-manipulators to move the mouse.
3. Non-contrast imaging protocol
- Measure the bladder wall thickness using the linear distance measurement tool and tracing the outside edge to inside edge of the bladder wall b-mode post acquisition.
- Measure the bladder 3D volume with the volumetric tool on the 3D mode acquisition by tracing the inside of the bladder walls to create a contour. Multiple contours are then generated through the thickness of the bladder to calculate volume.
4. Microbubble contrast resuspension/activation
- Activate the contrast agent (e.g., DEFINITY) by shaking in the vortex mixer for 45 s to encapsulate the microbubbles in solution. This step is critical for optimum contrast enhancement.
5. Catheter insertion
- With the mouse anesthetized and taped to the heated platform, expose the bladder with a midline incision using straight sharp/blunt scissors through the skin and abdominal wall.
- Insert a 27-gauge needle connected to a syringe by flexible polyethylene tubing (PE 50) into the bladder. Prefill the tubing with saline to ensure no air bubbles are injected into the bladder.
6. Contrast-enhanced imaging protocol
- To confirm needle placement, instill 10 µL of saline into the bladder while being observed via ultrasound.
- Replace the saline syringe with a syringe containing contrast to improve visualization of urethral walls and voiding events, because the urethra is normally collapsed. Once a complete long axis view of the urethra is obtained and an image saved, rotate the probe 90° to obtain a short axis view and an M-Mode image.
- Instill a bolus of microbubbles at 0.5 mL per 3 s into the bladder until a urination event occurs.
- During the first voiding event, measure the urethral diameter at the point of the Doppler velocity sample window using the linear distance tool and measuring edge to edge.
- With the urethra properly located, angle the probe in relation to the urethra to become more parallel to the urine flow.
- Instill a second bolus of microbubbles into the bladder, and measure the event velocity using the velocity time integral (VTI) tool.
- After data collection, euthanize the mouse with cervical dislocation.
7. Data Calculation and Analysis
- Select the VTI tool to measure velocity by tracing the recorded images.
- Measure the diameter of the urethra from the B-mode or M-mode image using the leading edge to leading edge convention.
- Calculate the cross-sectional area (CSA) using the following formula (CSA ) using image measurements obtained above.
- Calculate the void volume using the CSA of the urethra and multiplying that by the area under the Doppler trace (velocity time integral)(CSA x VTI = volume).
- Calculate the actual voided urine volume assuming one gram per cubic centimeter density.
Ultrasound can be used with or without contrast enhancement depending on the experimental design and endpoint measurement. Mice are anesthetized with isoflurane and shaved and all traces of hair removed with a depilatory cream. Anesthetized animals are placed on a heated platform with the ultrasound probe positioned along the long axis of the bladder (Figure 1).
Figure 2 shows representative ultrasound images of a mouse bladder acquired without contrast agent. The bladder wall is hyperechoic (white), and the bladder wall thickness is measured using a software measurement package. A bladder surface projection can be rendered to determine bladder volume, wall thickness, and wall volume (Table 1).
For contrast enhanced bladder and lower urinary tract imaging, a catheter must be inserted into the bladder and microbubbles injected. It is important to activate the microbubbles per manufacturer’s protocol for optimal imaging. Figure 3 shows a representative image of a microbubble-filled bladder. The bladder is hyperechoic (appears white in the image). A low frequency ultrasonic burst destroys the bubbles and the bladder becomes transiently hypoechoic (appears black), before the bubbles reform, confirming this structure as the bladder. During the destruction pulse, the transmit power goes to 100% and the transmit frequency drops to 10 MHz. A bolus of microbubbles (0.5 mL per 3 s) triggers a voiding event and makes it possible to visualize the urethra. The urethra is confirmed by applying a low frequency ultrasonic burst during the urination event and observing destruction and reformation of microbubbles. Several measurements can be made during the urination event. The urethral lumen diameter can be acquired pre- and post-urination along with the flow velocity of urine passing through that region of the urethra and total outflow (Figure 4). Using contrast imaging, measurements were made through the entire length of the urethra (Table 2). From these measurements, further calculations are made to examine urinary flow and bladder compliance (Table 3).
Figure 1. Ultrasound setup. (A) Overall imaging setup. (B) Positioning of mouse on platform. (C) Bladder is exposed and catheterized with the ultrasound probe lined up for long axis bladder imaging. (D) Ultrasound gel and probe placed on exposed, catheterized bladder for long and short axis for imaging. Please click here to view a larger version of this figure.
Figure 2. Non-contrast imaging of mouse bladder. (A) Image of mouse bladder without microbubbles. (B) Measurements of bladder wall thickness from b-mode bladder image. (C) 3D reconstruction of mouse bladder. (D) Surface area and shape extrapolated from 3D image for further analysis. Please click here to view a larger version of this figure.
Figure 3. Contrast-enhanced imaging of mouse bladder and urethra. (A) Full bladder with microbubbles. (B) Full bladder with microbubbles after destruction event. (C) Mouse urethra with microbubbles. (D) Mouse urethra with microbubbles after destruction event. Urethra outlined in red. Please click here to view a larger version of this figure.
Figure 4. Measurements taken from mouse urethra. (A) Urethral lumen diameters measured during a urinary event. (B) Urinary flow velocity through the penile urethra during voiding. Please click here to view a larger version of this figure.
|Trial||Bladder volume (mm3)||Bladder wall thickness (mm)||Bladder wall volume (mm3)|
Table 1. Non-invasive ultrasound measurements of the bladder. Bladder volume and bladder wall thickness measured by ultrasound.
|Measurement Type||Image Required||Location||Measurement|
|Bladder volume (mm3)||Bladder 3D Mode||Bladder||335|
|Bladder wall thickness (mm)||Bladder B-Mode||Distal to bladder neck
Proximal to bladder neck
|Urethra diameter (mm)*||Urethra B-Mode||Bladder neck
|Urinary event time (ms)*||Urethra PW mode||Bladder neck
|Acceleration time (ms)*||Urethra PW mode||Bladder neck
|Acceleration (mm/s2)*||Urethra PW mode||Bladder neck
|Velocity time integral (cm)*||Urethra PW mode||Bladder neck
|Mean velocity (mm/s)*||Urethra PW mode||Bladder neck
|Peak velocity (mm/s)*||Urethra PW mode||Bladder neck
|Mean gradient (mmHg)*||Urethra PW mode||Bladder neck
|Peak gradient (mmHg)*||Urethra PW mode||Bladder neck
|* Measurements captured during urinary event|
Table 2. Ultrasound measurements of the bladder and urethra. Bladder and urethra measurements made by ultrasound during urinary voids.
|Cross-sectional area (mm2)||CSA = πr2|
|Flow rate (mm3/s)||Flow rate = CSA x Mean Velocity|
|Estimated void volume (mL)||V = (Flow Rate/1000) x (Event Time in Seconds)|
|Volumetric stretch||Stretch = (Vafter-Vbefore)/Vbefore|
Table 3. Calculations using ultrasound measurements. Calculations and formulas applicable to ultrasound measurements to assess bladder function and urinary flow.
Current techniques for evaluating the lower urinary tract of rodents are limited by their ability to directly correlate changes in voiding physiology with changes in prostatic histology consequent to disease progression. Void spot assays and uroflowmetry can be used to assess spontaneous urination events in rodents, and these techniques can be used to evaluate changes over a period of time15,16,17. However, for both techniques, bladder fullness cannot be assessed prior to the start of the test. Additionally, changes in urination may occur due to behavior rather than a direct result of disease progression, making it difficult to determine the impact of disease on urination. Cystometry, another technique for evaluating bladder dysfunction in rodents, can provide an in vivo assessment of bladder function16. However, the dynamic effect of the rodent prostate on voiding function is not clear. Previous studies have documented mouse urethral histological changes in mice associated with altered voiding function12. However, these studies can only look at one discrete time point, and bladder, prostate, and urethral anatomy are not assessed at the same time as functional testing. Other methods of evaluating changes within the bladder (i.e., mass, volume) occur at time of euthanasia11,12, rendering it impossible to the observe evolution of a disease process over time. Because there is the risk of urination at time of sacrifice as well as the inability to regulate the water intake prior to sacrifice, the variability of measured bladder volume increases even within treatment groups. This paper describes the use of ultrasound to image the lower urinary tract of mice with or without a contrast agent. This technology allows for the visualization of changes in bladder size in an intact animal, as well as assessment of functional changes of bladder volume, bladder wall thickness, urethral lumen diameter, and velocity of contrast passing through the urethra. Specifically, contrast-enhanced ultrasound allows for the visualization of the urethral lumen, specifically in the prostatic region, during voiding in a fashion that pinpoints a region of potential dysfunction.
To ensure collection of consistent and accurate ultrasound data, it is crucial that a single trained sonographer collects and reads ultrasounds throughout the course of the study. For contrast enhanced imaging, it is important to activate the commercial microbubbles according to the manufacturer protocol. Activated microbubbles should be diluted with 0.9% saline solution. Undiluted microbubbles are so concentrated that they prevent ultrasound wave penetration and will shadow structures lying below them. Microbubble dilution also reduces experimental costs. Microbubble dilutions can be varied without negative experimental effects as needed by the user.
Evaluation of bladder volume and bladder wall thickness in an intact animal allows examination of disease progression and evaluation of treatment efficacy over time. Currently, treatments for BPH are administered in rodent models either as the disease is induced or at a time point previously determined to result in significant disease progression11,22,23. Efficacy of treatment is typically determined after a single, discrete period of time, despite the fact that biological variability may affect time required to respond to treatment. Using this novel technique, a rodent model for BPH can be evaluated from the induction of the disease phenotype through the entire treatment protocol.
A contrast agent enables the lower urinary tract to be visualized before, during, and after a urination event. We have previously examined urethral histology in a mouse model of BPH. We localized the prostatic urethra as the putative region giving rise to urinary dysfunction. This region contains more prostatic ducts, denser collagen, and a smaller lumen than in control mice12. Additionally, the BPH-susceptible mice demonstrate voiding dysfunction as measured by uroflowmetry and void spot assays. Using ultrasound with microbubbles, we can directly evaluate the region of the prostatic urethra to measure flow velocity, duration, and luminal diameter (Figure 3 and 4). By identifying the region where flow is impeded using ultrasound, that specific region can then be further evaluated histologically to determine the main component of dysfunction.
This technique is reproducible across mouse strains and across a range of mouse ages and treatment conditions. In addition to aged, male mice, this technique can be used to evaluate younger mice with metabolic abnormalities that could lead to BPH/LUTD. The technique can also be used to evaluate female mouse voiding and lower urinary tract function. Although the ultrasound protocol with contrast described here is a terminal procedure, we can perform a suprapubic cystostomy, creating the potential for non-terminal contrast imaging of the lower urinary tract24. Future experiments will optimize visualization of urinary tract functions to allow for repeated measures. Depending on the experimental questions, the techniques described here can be combined with other functional urinary testing techniques to gain more insight into disease progression and treatment efficacy.
The authors have nothing to disclose
We would like to thank Emily Ricke, Kristen Uchtmann, and the Ricke lab for their assistance with animal husbandry and feedback on this manuscript. We would like to thank the NIDDK and NIEHS for their financial support for these studies: U54 DK104310 (WAR, JAM, PCM, CMV, DEB), R01 ES001332 (WAR, CMV), K12 DK100022 (TTL, AR-A, DH). The content is the sole responsibility of the authors and does not represent the official views of the NIH.
|21mm Clear Tubing||Supera Anesthesia Innov||301-150|
|27 gauge needle||BD||Z192376|
|4 port Manifold||Supera Anesthesia Innov||RES536|
|DEFINITY||Lantheus Medical Imaging||DE4|
|F/AIR Canister||Supera Anesthesia Innov||80120|
|Graefe forceps (Serrated, Straight)||F.S.T.||11050-10|
|Inlet/Outlet Fittings||Supera Anesthesia Innov||VAP203/4|
|Isoflurane||Midwest Vet Supply||193.33161.3|
|Isoflurane Vaporizer||Supera Anesthesia Innov||VAP3000|
|MV707 probe||Fujifilm VisualSonics Inc|
|Oxygen Flowmeter||Supera Anesthesia Innov||OXY660|
|Polyethylene 50 tubing||BD||427516|
|Pressure Reg/Gauge||Supera Anesthesia Innov||OXY508|
|Rebreathing Circuits||Supera Anesthesia Innov||CIR529|
|Small Mice Nose Cone||Supera Anesthesia Inov||ACC526|
|Sterile saline||Midwest Vet Supply||193.74504.3||NaCl 0.9%, Injectable|
|Straight Sharp/Blunt Scissors||Fine Scientific Tools (F.S.T)||14054-13|
|Vevo 770||Fujifilm VisualSonics Inc|
|VIALMIX||Lantheus Medical Imaging||VMIX|
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