Doppler Ultrasonography for Live Imaging and Quantification of Ovarian Vascular Function in Mice

Abstract

To effectively assess ovarian vascular function and understand its role in ovarian physiology and pathology, non-invasive in vivo approaches are essential for capturing real-time changes without disrupting normal blood flow. Doppler ultrasonography is a well-established, non-invasive tool for assessing ovarian blood flow in larger species, but its application has been limited in mice, which are one of the most widely used animal models for research in ovarian biology, such as ovulation. Ovulation is a tightly regulated process that depends on coordinated follicular maturation stimulated by follicle-stimulating hormone, followed by a preovulatory luteinizing hormone (LH) surge that leads to rupture of the follicle wall and release of oocytes for fertilization. The LH surge also triggers a series of structural and functional changes in the ovarian vasculature (vascular remodeling), such as angiogenesis and constriction of capillaries at the follicular rupture site shortly before ovulation. In addition, a rapid increase in ovarian blood flow following the LH surge has been reported in multiple species but not in mice. This protocol utilizes Doppler ultrasonography to visualize the murine ovarian vasculature and quantify hemodynamic parameters. The protocol presented here reports detailed methods for hormone priming, anesthesia, positioning of the mouse, identification of the ovary and its vasculature using power and color Doppler, and measurement of blood flow velocity and resistance parameters. This method enables real-time, longitudinal assessment of ovarian vascular function in live mice, providing a powerful tool for studying ovarian vascular function in both physiological and pathological contexts.

Introduction

The ovary is a highly vascularized organ, and tightly regulated cyclical, structural, and functional changes in its vasculature (vascular remodeling) are essential for normal ovarian physiology, including follicle development, ovulation, and corpus luteum formation^{1 , 2 , 3 , 4} . Vascular remodeling encompasses a series of coordinated processes such as changes in vascular permeability, angiogenesis, vasodilation, vasoconstriction, and changes in blood flow (hemodynamics). These processes enable the ovary to rapidly adjust blood supply in response to hormonal cues across the estrous cycle^{1 , 2 , 3 , 5 , 6 , 7 , 8 , 9} . Disruptions in vascular remodeling and ovarian blood flow impair ovulation and are implicated in fertility disorders such as polycystic ovary syndrome^{2 , 3 , 10 , 11} . Obesity has similarly been shown to disrupt angiogenesis and to reduce the expression of vascular mediators such as endothelin-2^{12 , 13} . Additionally, ovarian hyperstimulation syndrome, a complication mostly associated with ovarian stimulation through gonadotropin treatment during in vitro fertilization, often presents increased stromal blood flow in the ovary, emphasizing the detrimental consequences of dysregulated vascular function¹⁴ . Collectively, these conditions highlight the central role of ovarian vascular remodeling in maintaining female fertility and underscore the importance of hemodynamics in supporting ovarian function.

Accurate evaluation of ovarian vascular function requires real-time measurement of hemodynamics without disrupting the physiological environment. In humans and large animal models such as cattle, sheep, and horses, Doppler ultrasonography is a widely used, non-invasive tool for assessing ovarian blood flow and vascular architecture^{15 , 16 , 17 , 18} . This technique enables dynamic analysis of ovarian vascularization and captures hormone-driven changes in blood flow, offering valuable insights into ovarian function and dysfunction^{15 , 16 , 17 , 18}.

Despite the extensive use of lab mice as a model for research in ovarian biology owing to their short reproductive cycle and genetic tractability, non-invasive, in vivo methods for studying ovarian blood flow in this model remain limited^{19 , 20}.

Existing approaches rely on highly invasive procedures that require externalization of the ovary (intravital microscopy), or on techniques such as the CLARITY approach or wholemount imaging that lack temporal resolution and disrupt physiological context^{1 , 2 , 3 , 4} . As the most used mammalian model for genetic manipulation and reproductive studies, the mouse presents an opportunity to explore molecular mechanisms of ovarian hemodynamics and evaluate targeted interventions. This highlights the need for non-invasive, longitudinal, and quantitative imaging approaches to monitor dynamic ovarian vascular changes in vivo.

The ovulatory process, a central event in female fertility, exemplifies the importance of ovarian vascular remodeling in female fertility. It begins with follicle-stimulating hormone (FSH)-induced follicular maturation and culminates in the LH surge that triggers follicular rupture and oocyte release¹ . Following stimulation with equine chorionic gonadotropin (eCG), which mimics the effects of FSH, the ovarian vascular network expands² . This vascular growth progresses outward from the ovarian medulla toward the cortex to support the developing follicles² . Shortly before ovulation, blood vessels extend into the granulosa cell layer, and localized vasoconstriction is observed at the follicular apex -- the site of eventual rupture and oocyte release³ . These coordinated vascular changes are critical for successful ovulation^{1 , 2 , 3} . Doppler studies in large animal models and humans have revealed changes in hemodynamics during the preovulatory period, including a rapid increase in ovarian perfusion and localized redistribution of blood flow within the preovulatory follicle, marked by increased flow at the base (closest to the ovarian stroma -- opposite to the apex) and decreased flow at the apex¹⁶ . The increase in velocity of ovarian blood flow immediately following the LH surge was associated with successful ovulation in women¹⁵ . However, no studies have yet longitudinally tracked these hemodynamic changes in the intact murine ovary, limiting our understanding of vascular remodeling in this widely used model.

To address this gap, we present a protocol for Doppler ultrasonography that enables non-invasive visualization of the murine ovary and its vasculature, along with quantification of ovarian hemodynamics, including velocity and resistance indices. Although the MS-700 probe used in this study has limited resolution (axial/lateral: 30/58 μm), which prevents detailed analysis of capillaries, it still allows clear visualization and quantification of the major ovarian blood vessels. The use of contrast agents in future applications may further enhance vascular visualization. Overall, this approach allows for repeated, longitudinal measurements in the same animal across defined time points, such as during the preovulatory period, while preserving physiological integrity, a key advantage over existing invasive techniques.

Protocol

All mice used were maintained in strict accordance with the National Institute of Health Guide for the Care and Use of Laboratory Animals, with ethical approval from the Institutional Animal Care and Use Committee at Cornell University. All animals used in this protocol were immature (21-23-day-old) C57BL6 mice.

1. Hormonal stimulation for superovulation

1. Inject 21-23-day-old mice weighing 10-12 g intraperitoneally with 5 IU of pregnant mare serum gonadotropin (PMSG) followed by 5 IU of human chorionic gonadotropin (hCG) 48 h later. Prepare each injection by diluting 50 μL of hormone (reconstituted from a 5000 IU stock vial) in 950 μL of sterile phosphate-buffered saline, and administer 0.1 mL per mouse.

2. Ultrasound machine turn-on sequence and transducer setup

1. Switch on the main power located on the back of the imaging system. After the system boots, toggle the computer standby switch on the left side of the cart to wake the monitor and computer.

2. Connect the desired transducer (MS-700 used here - 30-70 MHz) to the active transducer port on the imaging unit (rightmost port). Align the locking pin on the transducer connector with the notch in the port, then push the connector in fully and turn the locking lever to the vertical position to secure it.

3. Launch the analysis software by clicking on the software icon. Once the program opens and the transducer has been detected (automatically), select the ovary application package. Click New to create a new study and choose the appropriate name from the drop-down menu. In the Study Information window, enter the required fields: Study name; Series name; Application package: Ovary; Measurement package: Vascular.

3. Anesthesia, mouse preparation, and ultrasound stage setup

1. Turn on the physiological monitoring unit (temperature and heart rate). Ensure the platform is heated (37 °C).

2. Check that the isoflurane level is above the minimum fill line on the vaporizer reservoir. If necessary, add isoflurane using proper personal protective equipment. Open the oxygen tank valve slowly and completely. Set the oxygen flow rate at 1 liter per minute.

3. Adjust the stopcock on the y-piece tubing to allow oxygen and isoflurane to flow into the induction chamber. Ensure the stopcock leading to the nose cone on the mouse platform is closed during this step. Set the vaporizer dial to deliver 3-4% isoflurane (vol/vol) for induction.

4. Place the mouse into the induction chamber and close the lid securely.

   NOTE: The induction chamber is functionally airtight. Never leave an animal in a sealed chamber without active gas flow.

5. Prepare the imaging platform by applying a small amount of electrode cream to each of the electrodes to ensure proper contact for heart rate monitoring.

6. Once the animal is fully sedated, redirect gas flow by opening the stopcock to the nose cone on the imaging platform and closing the stopcock to the induction chamber.

7. Gently transfer the mouse onto the heated imaging platform. Place the mouse in the prone position (dorsal side facing upwards) (Figure 1A). Place the nose cone securely over the animal’s nose, ensuring both the nose and mouth are covered to maintain anesthesia. Tape each foot in the electrode cream onto the corresponding electrode pad for physiological monitoring. Apply ophthalmic ointment to prevent corneal drying during anesthesia.

8. Reduce the isoflurane concentration to 1.5-2.0% to maintain anesthesia by turning the knob to the left. Confirm adequate anesthesia depth using the toe-pinch reflex method (no withdrawal should be observed). Carefully monitor the animal’s vital signs every 5 min during the imaging session: (i) rate of breathing; (ii) rate of heartbeat; (iii) any movements that may indicate discomfort; and (iv) ensure that the imaging platform is heated to the correct temperature throughout.

9. Prepare the lower right quadrant of the back by shaving the area with an electric shaver. Shave from the right hindlimb up to the midline of the back, staying just lateral to the spine. Apply a thin layer of body hair remover cream to the previously shaved area for 30 s to 1 min to remove any remaining hair. Wipe off thoroughly with a damp gauze. Apply a generous, even layer of ultrasound gel to the shaved region to ensure optimal acoustic coupling.

10. Position the transducer probe in transverse mode relative to the mouse in the mechanical stand with the orientation notch facing to the left of the operator. Secure the probe in place using the clamp.

Figure 1: Mouse setup on imaging platform and diagram of the vessels supplying the ovary and the uterus.

(A) Representation of mouse setup on the ultrasound platform with the transducer placed at a 90°angle relative to the platform on the shaved lower-right quadrant of the mouse. (B) Graphical representation of blood vessels supplying the ovary and uterus. OA: Ovarian Artery; OV: Ovarian Vein; MV: Medullary Vessels; CV: Cortical Vessels; POAT: Periovarian adipose tissue; UA: Uterine Artery; UV: Uterine Vein. Please click here to view a larger version of this figure.

4. Identification of the ovary and ovarian blood vessels using Doppler ultrasonography

1. On the control panel, press the B-Mode key to activate the standard grayscale imaging window, which allows visualization of anatomical structures.

2. Gently lower the transducer onto the shaved area coated with ultrasound gel.

   1. Ensure that the imaging platform and mouse are positioned flat (parallel to the bench surface), and that the transducer is lowered perpendicular (90° angle - vertical) to the mouse and platform (Figure 1A).

   2. To locate the ovary, pull the skin towards the left, then position the transducer near the upper region of the shaved quadrant, closer to the midline of the back. Align the left edge of the probe adjacent to the spine, with the right edge extending slightly over the right lateral flank.

3. To identify the ovary, first detect the kidney, a large, light grey ovoid structure (5 mm × 6 mm) with visible cortex and medulla. Then push the imaging platform slightly forward (towards the head of the mouse), making the ovary and periovarian adipose tissue (POAT) appear.

   NOTE: The ovary is a small, circular, hypoechoic (dark gray) structure, approximately 1 x 2 mm (at preovulatory stage), located lateral to the spinal cord. The POAT appears hyperechoic (bright white).

4. To capture and store videos of the ovary in B mode, wait for the number of frames to increase (up to 100 frames), and press the Cine Store button on the keypad.

5. To confirm ovary identification and visualize blood vessels with high sensitivity to low-velocity flow, switch from B-Mode to power Doppler Mode by pressing the designated key on the control panel.

6. Adjust the following power Doppler settings for optimal visualization of ovarian vasculature by turning knobs on the keypad: Doppler Gain: 32-55 dB; Sensitivity: 5; Dynamic Range: 15 DR; Velocity 1 kHz.

   NOTE: The values can be viewed on the left-hand side panel and will change when the knobs are turned

7. In power Doppler Mode or color Doppler Mode (press the designated key on the keypad), identify the following vessels by moving the platform gently back and forth to visualize different aspects of the vasculature: (i) The ovarian artery (OA), branching from the abdominal aorta and coming from the direction of the POAT to enter the ovarian hilum; (ii) The medullary vessels (MV) within the central medulla; and (iii) The cortical vessels (CV) surrounding developing follicles in the outer cortex (Figure 1B).

   NOTE: Cortical vessels may not be consistently visible due to their small diameter and low flow velocity. Vessels at the base of the follicle are more easily detectable, while those at the apex are often not visible, particularly right before ovulation.

8. Press Cine store to capture videos of the ovary and its vasculature in power or color Doppler mode at any time to save the video currently displayed on the screen.

9. To distinguish between arterial and venous flow, press the color Doppler Mode key on the keypad. In this mode, flow direction is color-coded: red indicates flow toward the transducer, while blue indicates flow away from it. The vessel with the higher average velocity is the ovarian artery.

10. Changes in blood perfusion can be appreciated in color or power Doppler mode by visualizing the size of blood vessels and color intensity (yellow in power Doppler, red or blue in color Doppler).

5. Assessment of hemodynamics with color and power Doppler ultrasonography

1. While in either color Doppler or power Doppler mode, press the Pulsed Wave (PW) Doppler button on the ultrasound system keyboard. This brings up the sample volume gate (i.e., the two horizontal lines). Press the PW button a second time to activate the Doppler spectral waveform in the lower panel.

2. Position the sample gate (adjustable measurement tool - allows the user to define the area where velocities will be measured) within the center of the vessel lumen, ensuring alignment with the direction of blood flow. Adjust the insonation angle to be as parallel as possible to the vessel and maintain it at ≤60° to ensure accurate velocity calculations. Adjust the insonation angle by turning the corresponding angle knob.

3. Place the sample gate at the following positions for each vessel type. Ovarian artery: within the vessel just proximal to its entry point into the ovary. Medullary vessel: at the central branching point, where the MV branches into cortical vessels. This branching point appears as a pool of blood right after entrance into the ovary. Cortical vessel: within a vessel at the base of a growing follicle, when identifiable.

4. Once the sample gate is positioned at the desired location and a clear pulse waveform is visible, press Cine Store to record and save both the ultrasound image and the corresponding spectral Doppler waveform.

5. Once imaging is complete, gently remove the mouse from the imaging platform. Wipe off any ultrasound gel from the back and place the mouse on a heating pad until it wakes up fully.

6. Exporting and saving data

1. After completing all imaging and saving the desired still frames and cine loops using the Cine Store function, press the Study Management button located on the upper left side of the ultrasound system keyboard. This will open the Study Browser window, displaying a list of all saved images and cine loops from the current session.

2. In the Study Browser window, select the files to be saved.

   1. To save an entire series, select the corresponding series name -- this will automatically highlight all associated images and cine loops. Click the Copy to button to proceed.

   2. When prompted, click on the destination folder or external drive where the exported files should be saved. Confirm that the file format and destination path are correct before finalizing the export.

7. Turn off the sequence and clean up

NOTE: These steps should be completed immediately after imaging is concluded and all data have been saved and exported.

1. After exporting all necessary data, click the Shutdown icon in the Study Browser window of the analysis software application. This will safely power down both the acquisition computer and the control panel.

2. Allow the system cooling fans to run for 5-10 min. Once cooling is complete, switch off the main power using the toggle located on the back of the imaging cart.

3. Turn off all auxiliary equipment: oxygen supply, isoflurane vaporizer, and physiological monitoring unit.

4. Clean the ultrasound transducer: first wipe off any remaining ultrasound gel using lint-free wipes, then disinfect the surface with moist lint-free wipes or disinfectant wipes recommended for sensitive electronic equipment.

5. Clean the mouse platform and keyboard with disinfectant wipes. Do not use ethyl alcohol on the electrodes of the mouse platform, as it may damage sensitive components.

6. Wipe down the induction chamber with moist paper towels. Dispose of the paper towels in the biohazard waste container.

7. Dispose of all single-use materials (e.g., gloves, surgical tape, gauze, cotton-tipped applicators) in the designated biohazard container.

8. Measurement of hemodynamic parameters using Doppler pulse waveforms

1. In the analysis application, click copy from, select the appropriate name from the drop-down menu, navigate to where the study was saved, select it, and click okay to import it.

2. Open the desired PW Doppler image and click on the measurements icon in the bottom left corner to open the measurement panel.

3. Click the velocity measurement tool in the top left corner of the measurement panel to quantify the three highest peak systolic velocities (PSV) and end diastolic velocities (EDV) from the spectral Doppler waveform by clicking at the top of the PSV and EDV waves (the tool will automatically measure the size of the wave from the click point down to the x axis. Only include waveforms that display at least three distinct cardiac cycles with clear and consistent PSV and EDV peaks.

4. To calculate the average velocity, first compute the average of the three selected PSV values and the average of the three EDV values. Then calculate the mean velocity using the formula: Mean velocity = (Average PSV + Average EDV)/2.

5. For velocities below the detection threshold of 10 mm/s, assign a value of 5 mm/s (average velocity below threshold) or discard the measurement based on the research purpose.

6. To calculate the resistive index (RI), use the standard formula: RI = (PSV - EDV)/PSV. This index provides a measure of vascular resistance and should be computed using the averaged PSV and EDV values for accuracy.

7. To calculate the pulsatility index (PI), use the standard formula: PI = (PSV - EDV)/Average velocity

Representative Results

The objective of this protocol is to apply Doppler ultrasonography to visualize the murine ovary and its vasculature, assess changes in ovarian perfusion, and quantify ovarian hemodynamics.

Figure 2 shows representative images of the same ovary across multiple time points during the periovulatory period (12 h before and 12 h after ovulation) following hCG injection. Figure 2A shows B-mode images used to locate the ovary relative to POAT and the oviduct (step 4.3 in protocol). Figure 2B displays corresponding power Doppler images, which offer high sensitivity to blood flow and help confirm ovarian identification through the visualization of internal vascular structures (step 4.7). Power Doppler enables qualitative assessment of perfusion dynamics over time and identification of distinct vessel types, including the OA, MV, and CV. Increases in perfusion are reflected by enhanced signal intensity and vessel expansion. Although a standardized method for quantifying perfusion using Doppler is not yet established, power Doppler provides robust visual confirmation of changes in the structure and blood flow activity of ovarian blood vessels. Figure 2C features color Doppler images of the same ovary. This mode enables detection of flow direction and improves visualization of the OA because it allows for differentiation from the ovarian vein (see Step 4.9). Color Doppler also supports the qualitative assessment of changes in vessel size and serves as the preferred mode for PW Doppler velocity measurements due to improved localization of different vessel types and ovarian boundary compared to power Doppler. Using power Doppler, there was a visual increase of perfusion primarily in MV at 1 h post-hCG, followed by a return to the baseline at 8 h post-hCG and further decrease until immediately before ovulation at 12 h post-hCG. Collectively, these results demonstrate reliable ovarian identification and effective visualization of vascular dynamics using Power and Color Doppler imaging.

Figure 2: Identification of the ovary and visualization of the ovarian vasculature across the preovulatory and periovulatory periods using Doppler ultrasonography.

Representative images of the murine ovary at multiple time points following hCG injection, illustrating dynamic changes in blood perfusion across the preovulatory and periovulatory periods (A) B-mode images used for identifying the ovary and surrounding structures. Power (B) and color (C) Doppler images highlighting changes in ovarian perfusion and showing directional flow, respectively. n = 4. Circled area: ovary; OA: ovarian artery; MV: medullary vessels; CV: cortical vessels; POAT: periovarian adipose tissue; Ov.D: oviduct. Scale bar = 1 mm. The scale bar in the top image of each panel (A, B, C) can be applied to the other images in the panel. Please click here to view a larger version of this figure.

This protocol enables quantitative assessment of blood flow velocity using PW Doppler. Figure 3 presents representative images (Figure 3A,B) and corresponding PW waveforms (Figure 3C,D) from two mice: one with measurable OA flow (Figure 3A,C) and one with flow below the accurate detection threshold (Figure 3B,D). In both cases, the sample gate was placed over the OA, with the insonation angle aligned parallel to the vessel and maintained below 60 degrees to ensure accurate velocity estimation.

Figure 3: Detectable and undetectable blood flow in the ovarian artery.

(A-D) Representative color Doppler images (A,B) and corresponding pulse waveforms (C,D) from vessels with (A,C) and without (B,D) detectable flow. (A) Color Doppler image and (C) pulse waveform of detectable blood flow in OA at 1 h post-hCG. (B) Color Doppler image and (D) pulse waveform flow below the detection threshold in OA at 0 h post-hCG. n = 4. OA: ovarian artery; MV: medullary vessels; CV: cortical vessels; POAT: periovarian adipose tissue; Ov.D: oviduct. PSV: peak systolic velocity; EDV: end systolic velocity. Scale bar = 1 mm. Please click here to view a larger version of this figure.

In Figure 3C, the PW waveform exhibits at least three well-defined cycles with clearly distinguishable PSV and EDV values, each pair corresponding to a cardiac cycle. The three PSV and EDV values are used to calculate the average blood velocity. In contrast, Figure 3D shows a noisy trace lacking a discernible waveform, indicating flow below the reliable detection threshold of 10 mm/s. These low-signal traces typically occur in small vessels (e.g., CV) or in animals with poor hormonal response. In this example, the mouse in Figure 3B,D did not respond to hormonal stimulation and exhibited consistently low blood velocity at all time points (but not always below the detection threshold). This figure highlights the detection limits of the technique and provides guidance for identifying valid waveforms suitable for velocity quantification.

Figure 4 demonstrates the application of color Doppler ultrasonography for quantitative assessment of ovarian hemodynamics, including blood flow velocity and resistance index (RI). Figure 4A presents velocity measurements in the OA, MV, and CV from four mice throughout the preovulatory and periovulatory periods. There was an increase in velocity at 1 h post-hCG, followed by a return to baseline at 8 h post-hCG, which was maintained throughout the preovulatory period (12 h post-hCG) and into the periovulatory phase (24 h post-hCG), corroborating visual observations by power Doppler as shown in Figure 2. Mice 1-3 exhibited robust responses to hormonal stimulation, showing increased blood velocity at 1 h post-hCG in all three vessel types. In contrast, Mouse 4 did not respond to stimulation (no ovulation, lack of ovary enlargement, and minimal vascularization of the ovary and uterine horns observed during tissue collection), showing consistently low blood flow, with no velocity increase at 1 h post-hCG, particularly in the OA and MV. Mouse 4 serves as a representative example of the velocity range observed in mice that did not respond to hormonal stimulation. As expected, velocity in the CV was lower overall compared to OA and MV. In some cases, the velocity dropped below the detection threshold (~10 mm/s) in those vessels (Figure 4A). To indicate these instances (e.g., 8 h and 11 h in Mouse 4 and 24 h in Mouse 2 in the CV), a value of 5 mm/s was assigned to represent the mean of the velocities below the detectability limit (Figure 4A, right panel). Figure 4B shows RI measurements across the same mice, vessels, and timepoints. RI is an indirect measure of vascular resistance, which can influence perfusion and flow velocity. While these juvenile wild-type mice did not exhibit consistent RI changes during the preovulatory period, this parameter may be more informative in pathological models where vascular resistance is altered.

Figure 4: Longitudinal measurements of parameters of ovarian hemodynamics during the preovulatory and periovulatory periods.

(A-C) Longitudinal measurements of (A) blood velocity, (B) resistance index (RI), and (C) pulsatility index (PI) across the preovulatory and periovulatory timepoints in individual mice. Mice 1-3 exhibited a typical response to hormonal stimulation, while Mouse 4 showed no detectable response. Measurements were collected from three distinct vessel types: the ovarian artery, medullary vessels, and cortical vessels. n = 4. Please click here to view a larger version of this figure.

Figure 4C shows another measure of blood flow resistance: the pulsatility index (PI). PI compares the difference between peak and minimum flow velocity to the mean flow, which assesses flow variation and downstream resistance (after the point of measurement). As expected, PI showed the same overall pattern as the RI but on a different scale, as PI is normalized to mean rather than peak systolic velocity. No consistent changes in vascular resistance were observed across the preovulatory period. This likely reflects that resistance changes occur primarily at the microvascular level -- for example, within follicular capillaries that transiently constrict at the follicular apex to facilitate wall rupture -- rather than at the level of the entire ovary, which maintains blood flow to support developing follicles. Such microvascular changes cannot be detected by the Doppler ultrasonography equipment used in this study due to resolution limitations. Additionally, the inherently low flow velocity in the murine ovary, compared to larger organs like the kidney, reduces the difference between peak systolic and end-diastolic velocities, making subtle variations in flow resistance more difficult to detect.

Together, these results illustrate the capacity of Doppler ultrasonography to generate both qualitative and quantitative insights into ovarian vascular structure and function. This protocol offers a robust, non-invasive tool for tracking ovarian perfusion and hemodynamics and capturing inter-individual variability in response to hormonal stimulation.

Discussion

This study introduces a non-invasive method for visualizing and quantifying ovarian hemodynamics in vivo using Doppler ultrasonography. Unlike existing methods, this approach enables real-time assessment of blood flow without disrupting vascular function. Doppler ultrasonography is a widely used and effective tool for assessing ovarian morphology and vascular function in humans and large animal models^{15, 16, 17, 18} . Its application to murine ovarian physiology offers a valuable advancement in the study of ovarian vascular dynamics.

Doppler ultrasound provides a non-invasive and repeatable approach to assessing blood flow in mouse models, offering several advantages over traditional techniques. Conventional methods, such as radiolabeled microsphere injection, are highly invasive and terminal^{21 ,22 , 23} . This technique involves injecting microspheres into the left ventricle or an arterial catheter, where they become lodged in capillaries in proportion to local blood flow. A reference blood sample is collected simultaneously to calculate perfusion, after which the animal is euthanized for tissue radioactivity analysis^{21 ,22 , 23} . A similar technique can also be applied using fluorescent microspheres^{24 , 25} . Although sensitive and quantitative, this method does not allow for longitudinal measurements. More recently, intravital multiphoton imaging has enabled high-resolution visualization of blood flow within preovulatory follicles, including capillaries under 10 μm in diameter³ . However, this technique requires surgical externalization of the ovary, which is labor-intensive, may alter physiological blood flow, and has a limited time window for data collection before euthanasia of the experimental mouse (2-3 h under anesthesia). Finally, bi-directional arterial spin labeling magnetic resonance imaging (BD-ASL MRI) has been used to visualize perfusion in the murine placenta and may hold potential for application to the ovary²⁶ . However, this technique requires highly trained personnel to operate the MRI system, does not quantify blood velocity, and lacks the capacity for real-time imaging.

This protocol involves three critical steps: identifying the ovary, visualizing ovarian vasculature, and measuring hemodynamic parameters using PW Doppler. The following sections provide additional detail for each step and include troubleshooting recommendations to address potential challenges during imaging.

To optimize ovary identification, begin by gently pulling the skin toward the left of the mouse to expand the imaging area before lowering the transducer. If the resulting image lacks clarity, apply additional ultrasound gel to improve resolution. The ovary is most reliably located by first identifying the kidney, a large (~5 mm × 6 mm), easily distinguishable structure characterized by its size, unique vasculature (large cortical vessels branching outward), and distinct morphology (visible contrast between cortex and medulla). Once the kidney is visualized, move the imaging platform slightly forward (toward the mouse’s head) to bring the POAT into view. The POAT appears as a hyperechoic (bright white) region relative to surrounding tissues. The ovary lies just to the right of the POAT when the transducer’s orientation notch is positioned to the operator’s left. It can be identified as a ~1 mm × 2 mm circular structure in immature mice near the abdominal wall, with characteristic vascular branching within the medullary region. Unlike the ovary, adjacent tissues lack internal vascular signals. If the ovary is not clearly visible, slightly lift the transducer to reduce pressure on the abdomen, which may otherwise compress and obscure the organ.

The next critical step is to visualize ovarian vessels. Doppler ultrasonography allows for visualization and quantification of vessels at specific locations without interrupting signals from above regions. This spatial registration is achieved through pulse-delay encoding applied to each data column. The position of each signal is determined by the pulse delay and the speed of sound. Consequently, when the Doppler sample volume is manually positioned over a specific vessel, its location is distinguished from surrounding regions based on this pulse-delay encoding. The lateral and axial resolutions with the MS-700 probe are specified at 30 μm and 58 μm, respectively, indicating the accuracy of this positioning. Once the ovary is located, scan gently by moving the imaging platform forward and backward to visualize different vessel types, including the ovarian artery, medullary vessels, and cortical vessels. The OA should be visible either at the top or bottom of the POAT and will always be coming from the POAT into the ovary. The OA is the main vessel supplying the ovary, and Doppler measurements are obtained just before its entry into the ovary (as shown in Figure 1, where the OA is located immediately outside the ovary). Measurements of the MV were taken at the primary branch within the ovarian medulla. This location corresponds to that described by Feng et al., where vessel diameter was quantified in that study² . The CV velocity was measured at the distal end of the main vessel, where it terminates in the ovarian cortex-examples of cortical vessels can similarly be observed in Feng et al. Because flow velocity in cortical vessels is typically low, only a single branch is often visible, whereas multiple branches are reported when using methods with greater resolution² . When multiple cortical branches were visible, the branch exhibiting the highest velocity was recorded. In future applications, velocity measurements from multiple cortical branches could be acquired and averaged to provide a more comprehensive assessment. Using power Doppler to confirm the ovarian vessel structures can be beneficial, as it is more sensitive to low flow and allows for better visualization of vessels with slow flow. Acquire multiple images to ensure comprehensive vascular documentation by pressing Cine Store each time a new view of the vessels appears. Given the inherent variability in hemodynamic measurements, it is beneficial to have an adequate number of biological replicates combined with outlier analysis to improve reliability. Alternatively, two independent operators can repeat experiments separately to identify consistent patterns of hemodynamics.

To ensure the most accurate velocity measurements are recorded, position the sample gate at the center of the target vessel. Adjust the insonation angle dial to align the angle parallel to the vessel, keeping it below 60 degrees. When the PW signal appears, use the velocity scale dial to produce an interpretable waveform, raising the scale for high-velocity flow and lowering it for slower flow. Record measurements at multiple sites along the vessel to calculate an average velocity. The minimum reliable threshold is 10 mm/s; vessels with lower flow may produce noisy, non-waveform signals. In addition to velocity data, color and power Doppler modes can be used to assess regional perfusion patterns within the ovary.

While Doppler ultrasonography offers key advantages, such as real-time visualization and quantification of blood flow without physiological disruption, it also has limitations. The primary limitation of this technique is its spatial resolution, which remains low relative to the small size of the ovary and its intricate vasculature, as well as the relatively slow rate of blood flow within these vessels. Using the ultrasound system, two probes are commonly used to visualize the murine ovary. The MS-700 probe, used in this protocol, operates at 30-70 MHz, with a maximum imaging depth of 10 mm and axial/lateral resolution of 30/58 μm at the center frequency. This probe is ideal for imaging juvenile mouse ovaries, as demonstrated here. Alternatively, the MS-550D probe (22-55 MHz) provides a greater imaging depth of up to 15 mm, with axial/lateral resolution of 40/80 μm, providing better images in adult mice due to increased penetration. Regardless of the probe selected, vessel structures smaller than 3 μm in diameter or located within 3 μm of each other cannot be reliably resolved due to resolution limits.

The diameter of the OA increases with hormonal stimulation, from approximately 35 μm before priming to 130 μm 48 h after eCG injection^{2 , 27} . This suggests that while the OA can be clearly visualized following stimulation, it may be difficult to detect under basal conditions. Additionally, the limited resolution of ultrasound prevents the detection of small changes in vessel diameter. For example, during the preovulatory period, the OA decreases in diameter by approximately 5 μm within the first 6 h following hCG injection, a change too subtle to be reliably detected by the method described in this protocol^{2 , 27} . Similarly, capillaries surrounding growing and preovulatory follicles are often below the resolution threshold and may not be accurately visualized. At the follicular apex, where ovulation occurs, capillaries measure as little as 10 μm and decrease further in size just before ovulation³, rendering them below the detection threshold of this imaging system.

In addition to the limitations imposed by spatial resolution, it remains challenging to clearly distinguish preovulatory follicles and CL. Large antral and preovulatory follicles are theoretically resolvable and can occasionally be observed; however, the overall echotexture of the mouse ovary and the limited contrast between follicular fluid and surrounding stroma make them difficult to identify reliably. Similarly, CL cannot be consistently distinguished from the surrounding tissue due to its heterogeneous echogenicity, at least at the early luteal phase (24 h post-hCG) when our measurements were taken. Consequently, this limitation prevents clear attribution of blood flow signals to specific anatomical structures (follicles versus CL). Although blood flow to specific structures is difficult to visualize, the precisely timed imaging following ovulation stimulation used in this protocol enables comprehensive quantification of ovarian hemodynamics in large vessels throughout the preovulatory to periovulatory period. Additionally, 3-dimensional (3D) Doppler ultrasound imaging could be applied here to obtain a more comprehensive view of the ovary, potentially improving the characterization of morphological structures such as preovulatory follicles and CL.

Although Doppler ultrasonography offers lower spatial resolution than histological methods, it provides superior temporal resolution, is non-invasive, and enables real-time assessment of dynamic vascular function in vivo, making it a valuable tool for studying ovarian physiology in the mouse model. Recent advances have further improved the resolution of Doppler imaging through the use of contrast agents, advanced signal processing techniques such as Doppler slicing, contrast-free super-resolution power Doppler (CS-PD), and ultrasound localization microscopy (ULM)^{28 , 29 , 30 , 31} . Several approaches have been employed to assess organ perfusion using Doppler ultrasonography^{32 , 33} . Some, such as the method used in this study, rely on measurements of the RI, whereas others estimate fractional moving blood volume (the percentage of tissue volume containing moving blood) from 3D Doppler imaging^{32 , 33} . In the current study, pixel-intensity measurements of ovarian perfusion were excluded because they were highly sensitive to motion artifacts (such as breathing rate) and the size of the region of interest (ovary); therefore did not reliably reflect physiological changes in perfusion. Future application of 3D imaging could provide a more comprehensive assessment of tissue perfusion. Overall, while these advanced techniques have not yet been applied to murine ovarian imaging, they hold significant potential for enhancing the accurate identification and measurement of ovarian blood vessels.

The application of Doppler ultrasonography, a well-established method for visualizing perfusion and quantifying hemodynamic changes, offers significant potential for advancing the understanding of ovarian physiology in the mouse. This technique enables both visual and quantitative assessment of ovarian hemodynamics in vivo and in real time. As vascular remodeling is a key component of follicle development and ovulation, the ability to repeatedly assess these processes without disrupting blood flow represents a major advantage over previous methods used in mice. This makes Doppler ultrasonography a particularly powerful tool for investigating murine ovarian vascular function. Future applications of this technique include tracking dynamic changes in blood flow across the estrous cycle and evaluating vascular disruptions in conditions such as PCOS and OHSS^{11 , 14} . The mouse’s unique suitability for genetic manipulation enhances the value of this method, enabling researchers to assess how specific molecular regulators affect ovarian perfusion and vascular dynamics in vivo. In addition to assessing blood flow, Doppler ultrasonography allows for the visualization of structural abnormalities such as cysts. To enhance the detection of ovarian morphological features (cysts, follicles, CL), future applications of this protocol could include the use of 3D imaging along with contrast agents. Integrating 3D Doppler imaging with the approach described here would potentially allow for a more complete and sensitive assessment of ovarian structures and ovarian blood flow dynamics, thereby enhancing our understanding of vascular remodeling and hemodynamic changes during the preovulatory period. Emerging techniques, including the use of contrast agents and CS-PD, offer the potential to enhance spatial resolution and enable more accurate measurement of blood vessel diameter, providing valuable insights into ovarian vascular remodeling. Furthermore, these advanced methods (ULM with contrast agents) have been successfully applied to quantify organ perfusion in other tissues. If adapted for ovarian imaging, they could offer an additional quantitative measure of hemodynamics and tissue function³¹ . As a clinically relevant, non-invasive, and evolving technique, Doppler ultrasonography also offers strong translational potential for bridging animal studies and human reproductive medicine. Together, these strengths position the application of Doppler ultrasonography to the mouse model as a foundational tool for advancing both basic and translational ovarian research.