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

Summary

Changes in ovarian blood flow during the preovulatory period are essential for ovulation. Doppler ultrasonography has been used in humans and large animals to assess ovarian hemodynamics and perfusion. A protocol is presented for applying Doppler ultrasonography to evaluate ovarian hemodynamics during the preovulatory and periovulatory phases 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 whole-mount 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.

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

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 PO...

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,

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3M Surgical tape	VWR International LLC	MMM1534-3
Aquasonic Clear Ultrasound Transmission Gel	Parker	PKR-03-08
Hair Clipper	Skull Shaver LLC	BBT 2020
Human Chorionic Gonadotropin	Sigma Aldrich	9002-61-3
Nair Body Cream hair remover	Nair		Hair removal
Optixcare Eye Lube Plus	Aventix	OPX-4252
Parker Signacreme Electrode Cream	Parker	PKR-17-05-CS
Pregnant Mare Serum Gonadotropin	Creative Enzymes	NATE-0969
VisualSonics Vevo 2100 Imaging system	VisualSonics Inc.