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Cancer Research

Non-Invasive Ultrasound Assessment of Endometrial Cancer Progression in Pax8-Directed Deletion of the Tumor Suppressors Arid1a and Pten in Mice

Published: February 17, 2023 doi: 10.3791/64732

Abstract

Uterine cancers can be studied in mice due to the ease of handling and genetic manipulation in these models. However, these studies are often limited to assessing pathology post-mortem in animals euthanized at multiple time points in different cohorts, which increases the number of mice needed for a study. Imaging mice in longitudinal studies can track the progression of disease in individual animals, reducing the number of mice needed. Advances in ultrasound technology have allowed for the detection of micrometer-level changes in tissues. Ultrasound has been used to study follicle maturation in ovaries and xenograft growth but has not been applied to morphological changes in the mouse uterus. This protocol examines the juxtaposition of pathology with in vivo imaging comparisons in an induced endometrial cancer mouse model. The features observed by ultrasound were consistent with the degree of change seen by gross pathology and histology. Ultrasound was found to be highly predictive of the observed pathology, supporting the incorporation of ultrasonography into longitudinal studies of uterine diseases such as cancer in mice.

Introduction

Mice remain one of the most important animal models for reproductive disorders1,2,3. There are several genetically modified or induced rodent models of ovarian and uterine cancers. These studies typically rely on multiple cohorts euthanized at different time points to capture longitudinal trends in morphologic and pathologic changes. This prevents the ability to acquire continuous data on cancer development in an individual mouse. Additionally, without knowing the individual mouse disease progression state, intervention studies are based on predetermined time points and averaged findings of previous cohorts rather than individual thresholds for the detection of progression in a specific animal4,5. Therefore, imaging approaches that allow for longitudinal assessment in live animals are needed to facilitate preclinical models for testing new drugs or compounds and accelerate the understanding of pathobiology while also increasing the rigor and reproducibility6.

Ultrasound imaging (US) is an appealing method for the longitudinal monitoring of mouse uterine cancer progression because it is relatively facile and inexpensive compared to other imaging methods, is easy to perform, and can have remarkable resolution6,7. This non-invasive modality can capture features to the micron scale in awake mice or with mice under brief sedation using a 5-10 min exam. Ultrasound microscopy has been validated as a method to measure mouse ovarian follicle development 8 and the growth of implanted or induced neoplasia9,10,11. High-frequency US has also been used for percutaneous intrauterine injections12 and observing rat uterine change over the estrus cycle13. High-frequency US can be used with mice held on specialized stationary platforms using a rail system to hold the transducer/probe to capture high-resolution images with standardized position and pressure; however, this equipment is not available at all institutions. Hand-held transducer scanning methods can be adopted with less dedicated equipment and used for both clinical diagnostics and research applications in mice.

The question remains as to whether US imaging with hand-held, high-frequency probes could be used to monitor uterine cancer development over multiple weeks. Similar to the intestines, the rodent uterus is a thin-walled, slender structure that is very mobile within the abdomen and is contiguous through multiple tissue depths, making imaging more challenging than with relatively immobile organs such as the kidneys. This study sought to establish the correlation between tissues observed by ultrasound and histopathology, define landmarks for locating the mouse uterus, and determine the feasibility of the longitudinal assessment of endometrial cancer. This study presents data showing a qualitative correspondence between the appearance of uteri imaged by US and histopathology, as well as serial imaging of mice over several weeks. These results indicate that hand-held US can be used to monitor endometrial cancer development in mice, thus creating an opportunity for collecting individual mouse longitudinal data to study uterine cancer without the need for dedicated equipment.

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Protocol

All procedures and experiments involving mice were performed according to protocols approved by Johns Hopkins Animal Care and Use Committee. For all procedures, appropriate PPE was worn, including gloves and disposable isolation gowns. Precautions were taken when handling sharps, which were properly disposed of in red box sharps containers immediately after use. See the Table of Materials for details about all the materials and equipment used in this protocol.

1. Induction of endometrial cancer in iPAD (inducible Pten, Arid1a double deletion) mice with doxycycline

  1. Maintain 10 Pax8-Cre-Arid1a-Pten double deletion (iPAD) transgenic mice (Figure 1) on a mixed genetic background (129S, BALB/C, C57BL/6), as previously described14.
  2. Collect baseline ultrasound (2D) images of the ovaries, oviduct, and uterus of each mouse before doxycycline treatment.
  3. Provide exclusively doxycycline-containing mouse chow diets (doxycycline hyclate at 625 mg/kg of feed) to the female iPAD mice for a minimum of 2 weeks starting at 7-8 weeks of age to induce gene deletion.

2. Equipment setup

  1. Turn on the heating pad, and cover with a clean absorbent pad (target temperature: 38 °C).
  2. Confirm that the isoflurane vaporizer and O2 tank are adequately filled. Refill and replace if the contents are low.
  3. Connect the induction chamber, nose cone, and scavenging system to the vaporizer.
  4. Set up the ultrasound machine.
    1. Select a transducer (probe) with a range of 32-56 MHz or up to 70 MHz for imaging of the uterus or ovary, respectively.
    2. Attach the probe, and power on the machine.
    3. After system boot-up, use the control panel to log in with the user credentials and access the home screen.
    4. From the home screen, go to the Applications tab, and select Mouse (small) abdomen mode.
    5. Click on Scan to return to the home screen, and wait for a live image to be displayed.
    6. Select B-Mode from the options on the left tool bar.
    7. Click on More Controls to view additional tools for image refinement, such as image gain and depth, or to adjust the clip acquisition settings, such as the number of frames per second.
    8. Once the image settings are selected, return to the home screen by clicking on Scan.
  5. Turn on the O2 tank, direct the flow to the induction chamber, and set the flow rate to 1 L/min.

3. Preparation of mice for ultrasound screening, including hair removal

  1. Place a mouse in the induction chamber. Set the isoflurane vaporizer to 2%-3% vol/vol for the induction of anesthesia.
  2. Determine the appropriate depth of anesthesia by a lack of response to toe pinch and a respiratory rate around 1-2 breaths/s.
  3. Remove the fur from the dorsum and ventrum between the last rib and pelvis with appropriately sized clippers.
  4. Apply a thin layer of depilatory cream to the ventral and dorsal regions to be imaged (if needed).
  5. Place the mouse back in the induction chamber for approximately 3-5 min to maintain the appropriate depth of anesthesia while the depilatory cream works to remove the hair. After ≤4 minutes, gently wipe away the cream with a clean moist paper towel.
    ​NOTE: Longer exposure to depilatory cream is irritating and may cause skin lesions.

4. Intraperitoneal injection of fluid to increase the contrast between organs

  1. Warm a 3-10 mL syringe filled with sterile isotonic fluid solution (e.g., sterile 0.9% NaCl or Lactated Ringers Solution) to 35-40 °C by placing it between a heating pad and an absorbent pad for several minutes. Place a bottle of ultrasound gel on the heating pad if the machine does not have a warmer.
  2. For a 20-25 g mouse, inject 1-2 mL of solution into the peritoneal cavity.
    1. Secure the mouse by the scruff in one hand, exposing the ventrum.
    2. Hold the mouse at a ~20° angle, with the nose pointed to the floor to direct the organs cranially due to gravity.
    3. Using a small gauge needle (25 G, 5/8 in length, tuberculin syringe), puncture through the skin and abdominal wall of the caudal right quadrant of the abdomen.
    4. Before the injection of fluids, to avoid injection into the vasculature or the GI tract, pull back with minimal pressure. If blood or other material enters the syringe, remove the needle. Use a new needle and syringe, and try again at a slightly different position.
  3. If the mouse wakes up during the injections, place it back in the small induction chamber for anesthesia with 2%-3% vol/vol isoflurane.

5. Ultrasound imaging from a dorsal approach

  1. Position the mouse in ventral recumbency on the absorbent pad over a heating pad (Figure 2A-C).
  2. Place a rodent nose cone securely over the mouse's nose and muzzle. Maintain the anesthetic depth with isoflurane delivered through the nose cone at 1%-2% vol/vol in 100% O2.
  3. Monitor the mouse for a regular respiratory rate (1-2/s) and a lack of toe pinch response to indicate if the anesthesia needs to be adjusted.
  4. Place a small amount (~0.5-1 mL) of prewarmed ultrasound gel abaxial (lateral) to the spine on either side of the anesthetized mouse, between the last rib and pelvis.
  5. Put a small amount of gel on the ultrasound probe.
  6. Place the probe parallel to the vertebrae with the front of the probe on the cranial side. An indicator mark is present on the probe head to indicate the proper probe orientation. Record the day, time, animal ID, probe orientation, and animal side (right, left, dorsal, ventral) for each new set of images being collected.
  7. With a mouse in ventral recumbency (dorsum skin touching the probe), slowly scan the area for the kidney landmark (Figure 2B and Figure 3). With the kidney in view, pull the probe caudal to find the ovary-a slightly hyperechoic oval to round structure (Figure 4A, B) within a very hyperechoic ovarian fat pad that is bordered cranio-ventrally by the kidney and dorso-laterally by the dorsal abdominal wall.
    NOTE: Pressure caudal and lateral to the ovary can direct the ovary closer to the abdominal wall and away from the loops of the intestine. The ovary is anatomically positioned up against the dorsal abdominal wall, just ventral and lateral to the epaxial muscles and caudal to the kidney.
  8. Adjust the signal gain using the slider at the bottom of the control screen to improve the image contrast.
  9. To improve the imaging of the kidney, apply pressure with a finger to the contralateral abdomen. Vary the pressure and angle from directly parallel to the spine to ~20° ventral.
  10. Once the organ of interest is in view, collect a video by clicking on Save Clip or Start Recording and then Stop Recording when done to retain images at a preset number of frames.
  11. Save single frames from either a live image or recording with the Save Frame button.
  12. To image the uterus, pull the probe caudally until the ovary is in the most cranial aspect of the field of view. Vary the probe pressure and angle until the uterus is in view.
  13. Repeat video and frame collection for each organ of interest.
  14. Find the uterus running longitudinal along the dorsal abdominal wall with the lateral leg musculature also in view (Figure 4B).
    NOTE: The uterus size and lumen diameter may vary with the phase of estrus and disease state.
  15. Monitor the tissue for peristaltic motion to differentiate the intestinal loops from the uterine stationary horns.

6. Collect images from a ventral approach

  1. Place the mouse in dorsal recumbency, and check that the eye lubrication is sufficient and the muzzle is securely in the nose cone (Figure 2A).
  2. Apply a small amount (~0.5-1 mL) of prewarmed ultrasound gel to the ventral abdomen, and apply the probe at the midline just cranial of the pubis to locate the bladder as a hypoechoic landmark (Figure 5).
    NOTE: If the bladder is too large and obscures the uterus imaging, gentle pressure can be placed on the lower abdomen to express urine.
  3. Pull the probe lateral to the bladder to find the uterine horns. Apply light digital pressure from either or both sides of the mouse to bring the horns into the field of view. Hold the probe perpendicular to the mouse, and scan both sides of the abdomen to capture transverse views (cross-sections) of both horns. Rotate the probe to capture sagittal views.
  4. After the ultrasound, wipe the mouse clean of gel with a paper towel, and return it to its cage to recover. Mice are fully awake in 2-5 min. Once it is fully awake and ambulatory, return the mouse to the animal room.
    NOTE: A heating pad on a low heat can be placed under the cage to warm the cage for recovery.
  5. At the experimental or humane endpoint, euthanize the mouse. Ideally, euthanize the mouse in the home cage to reduce stress; alternatively, place the mouse in a clean chamber. Deliver pressurized CO2 at a displacement rate of 10%-30% of the chamber volume per minute. After approximately 5 min of no visible respiration, verify death by cervical dislocation. Proceed with abdominal necropsy for tumor harvest.

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Representative Results

Pax8-Cre-Arid1a-Pten double deletion (iPAD) transgenic mice were maintained on a mixed genetic background (129S, BALB/C, C57BL/6), as previously described14. The mice were all fed a doxycycline feed for 2 weeks to induce Cre recombinase. In previous work by our group, doxycycline was dosed by gavage14; however, in this current study, the doxycycline feed induction method worked efficiently and reduced the stress of gavage for the mice. It is important to check that the doxycycline administration method is adequate to induce the Cre recombinase expression needed to excise the DNA for tumor suppressors (Arid1a and Pten) in Pax8-expressing cells, such as the uterus. Immunohistochemistry with antibodies directed against ARID1a and pTEN14 was used to check if adequate tumor suppressor protein loss was accomplished in the uterine epithelial cells compared to the neighboring stromal tissue, as seen in Figure 6. The schematic of the mouse model and tumor suppressor expression regulation is presented in Figure 1.

After 2 weeks of doxycycline, the uterus did not change significantly in size compared to controls or the same mouse imaged before doxycycline administration. Yet, over the next 2-6 weeks post doxycycline, ultrasound showed that the mice developed a range of uterine pathologies, beginning with hyperplasia and progressing to adenocarcinoma. Figure 7A,B depicts US images of a mouse uterus with early pathology changes seen after 3 weeks of doxycycline treatment. Figure 7C,D shows the gross pathology and histopathology of the same uterus imaged in Figure 7A,B after the mouse was euthanized immediately after imaging at 3 weeks. The smaller (but dilated) thin-walled uterus and early endometrial hyperplasia with preservation of the distinct lumen (central hyperechoic uterine fluid) can be observed. In mice euthanized at later time points, the uterus was much larger, had a smaller lumen, and had a thickened wall (Figure 8 and Figure 9). Examples of dilated lumens and subtle changes in the hyperplastic endometrium are compared to dense tissue and nodular cancer growth in the uterus in Figure 8 and Figure 9. The ultrasound and gross images were highly consistent with each other for all the represented mice. Figure 9 presents multiple uterus images from the same mouse in a time course study and demonstrates the morphological changes observed over 6 weeks as adenocarcinoma developed. Figure 5 presents a ventral approach with cross-sections of the uterine horns adjacent to the hypoechoic bladder.

To enhance the contrast in the abdomen, injecting saline into the abdominal cavity directly before imaging improved the visualization of the abdominal organs, including the uterus and ovaries. Figure 3 shows ultrasound comparisons from the same mouse before and after the injection of 1 mL of saline. With saline, there was an increased hypoechoic space (black) between the organs that allowed for the kidney, fat pad, ovary, uterine horns, intestines, and spleen to be more readily individualized from each other. In all the other figures presented here, saline injections were done before imaging to improve the image clarity.

Taken together, this protocol shows that this inducible mouse model develops adenocarcinoma over a 6 week period and can be monitored with ultrasound to establish a baseline and observe morphological changes in the uterus as the cancer develops. Here, it is demonstrated that non-invasive ultrasound allows for the uterine horns in a mouse to be serially assessed over multiple weeks with images that are predictive of the actual uterine pathological changes.

Figure 1
Figure 1: Strategy for the generation of the Pax8-Cre inducible double KO transgenic mice. (A) The constitutive production of a reverse tetracycline-controlled transactivator (rtTA) in Pax8-expressing uterine epithelial cells results in the inducible activation of Cre recombinase in the presence of tetracycline, or its analog doxycycline, by binding to tetracycline response elements (TRE). Flox sites located upstream and downstream of exon 8 (Arid1a) and exon 5 (Pten) are targeted by Cre recombinase. Cre-targeted deletion of exon 8 in Arid1a results in a frameshift mutation and an early stop codon (p.Gly809Hisfs*6). Cre-targeted deletion of exon 5 in Pten results in a frameshift mutation (p.Val85Glyfs*14). (B) The Pax8-rtTA mouse expresses a reverse tetracycline-controlled transactivator (rtTA) under the control of the Pax8 promoter. This mouse is crossed with the TetO-Cre mouse that expresses Cre recombinase in a tetracycline-dependent manner. The progeny from this crossing express a tetracycline (or doxycycline)-activated Cre recombinase specific to Pax8-expressing uterine epithelial cells (Pax8-Cre transactivator). (C) Arid1aflox/flox mice on the 129S1 background were crossed with Ptenflox/flox on the BALB/C background (Strain C;129S4-Ptentm1Hwu/J) to generate the Arid1aflox/flox/Ptenflox/flox transresponder mouse. (D) The Pax8-Cre transactivator mouse is crossed with an Arid1aflox/flox/Ptenflox/flox Transresponder mouse. Progeny from this crossing result in a transgenic murine model with a doxycycline inducible Arid1a and Pten deletion (iPAD) specific to uterine epithelial cells. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Imaging procedure. (A) An anesthetized mouse with the abdominal and dorsal fur removed with a clipper and depilatory cream. The mouse is maintained on inhalant isoflurane anesthesia via a nose cone and placed on a heat pad covered with an absorbent pad. (B) A small amount of prewarmed ultrasound gel is placed over the region of interest on the mouse and on the probe. Here, the mouse is in ventral recumbency, and the probe is correctly placed to locate the kidney, the anatomical landmark directly cranial to the ovary. (C) In dorsal recumbency, the probe can be manually manipulated to first find landmarks such as the bladder in order to find the approximate location of the uterine horns. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Improvement of visibility and organ definition by the injection of saline. (A) Image of a kidney and loops of the GI tract (bottom left) before the IP injection of 1.5 mL of warmed, sterile saline. The left side of the image is cranial to the kidney. (B) Image of the kidney after the saline injection, showing increased (hypoechoic space, black) between the organs. For both images, in terms of the orientation, the mouse is dorsal surface up, and cranial to caudal is left to right. (C,D) The highlighted red section shows kidney landmarks in the 2D images presented in A and B. The images were taken using an MS550 transducer. Scale bars = 2 mm in each image. Abbreviations: GI tract = gastrointestinal tract; IP = intraperitoneal. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Reproductive tract with landmarks. (A) The ovary (arrow) is located immediately caudal to caudal pole of the kidney. (B) The uterine horn (narrow arrow) is deep and begins cranial to the dorsal muscles of the proximal hindlimb (arrowhead), with the ovary also in view (bigger arrow). Scale bars = 2 mm. For all the images, in terms of orientation, the mouse is dorsal surface up, and cranial to caudal is left to right. The images were taken using an MS550 transducer. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Uterine horns and bladder imaged from dorsal recumbency. Ultrasound images from two mice before cancer induction. (A) The bladder lumen is visible as a hypoechoic, ovoid structure with a hyperechoic mucosa. Both uterine horns are visible together, with the boundaries of each traced in light blue. (B) The same view in another mouse with a much fuller bladder. For both images, in terms of orientation, the mouse is ventral surface up, and the image left is the mouse left. The boundaries of each uterine horn are drawn in light blue, with areas given in the neighboring text boxes (MS550 transducer). Scale bars = 3 mm. Please click here to view a larger version of this figure.

Figure 6
Figure 6: Examples of H&E and immunohistochemistry of sections of uterus at 6 weeks post induction in an iPAD mouse. (A) Representative H&E of iPAD mouse uterine tissue. (B) Representative ARID1a immunohistochemistry. The staining shows a loss of Arid1a specific to the nuclei of Pax8-expressing uterine epithelial cells (black arrow), with the retention of Arid1a in the nuclei of the uterine stromal cells (white arrow). (C) Representative pTEN immunohistochemistry. The staining shows a loss of pTEN in the cytoplasm of Pax 8-expressing uterine epithelial cells (black arrow), with the retention of pTEN in the cytoplasm of the uterine stromal cells (white arrow). Detailed IHC methods have been previously published14. Scale bars = 100 µm. Abbreviations: iPAD = inducible Pten, Arid1a double deletion; H&E = hematoxylin and eosin. Please click here to view a larger version of this figure.

Figure 7
Figure 7: Examples of ultrasound images from mice with early changes in pathology. (A,B) Antemortem ultrasound image of a uterus filled with uterine fluid that is naturally hyperechoic The narrow arrows indicate the hypoechoic, dilated lumen of the uterus. The arrowhead indicates the thickened area of the uterine wall. (C) Gross image of the entire uterus from the mouse imaged in B. The white asterisk is roughly at the same location as that imaged in A. (D) In this same mouse, the uterus was processed and stained with H&E. The histology section shows thickening of the uterine wall due to endometrial hyperplasia; the example region is indicated with an arrowhead. For all the US images, in terms of orientation, the mouse is dorsal surface up, and cranial to caudal is left to right (MS550 transducer). Scale bars = (A,B) 3 mm; (D) 500 µm. Abbreviations: US = ultrasound; H&E = hematoxylin and eosin. Please click here to view a larger version of this figure.

Figure 8
Figure 8: Diversity of uterine pathology visible by ultrasound. (A) A uterus with hyperplasia and fluid distension causing luminal dilation to a diameter of ~2.5 mm. (B) A second uterus with hyperplasia that is distended to a greater degree. (C) A uterus that is both dilated and filled with a soft tissue mass. Scale bars = 3 mm. For all the images, the arrows are to the ventral side of the uterine horn, and in terms of the image orientation, the mouse is dorsal surface up, and cranial to caudal is left to right (MS550 transducer). Please click here to view a larger version of this figure.

Figure 9
Figure 9: Changes in the neoplastic development of the uterus over time in one mouse. (A) Ultrasound image of an iPAD mouse showing the kidney (wider arrow), ovary (caudal to kidney), and uterus (narrow arrow), with the leg muscle also visible (arrowhead), captured at three time points: after the mouse was started on a doxycycline diet (week 0), and then again after 5 weeks and 6 weeks. Doxycycline induced Cre expression with tumor suppressor loss, which promotes endometrial cancer over time. (B) The same ultrasound images as A but with the uterine horn highlighted in red. (C) Gross images of the uterus in situ (narrow arrow) from the 6 week time point; the tissue is viewed from the ventral surface. (D) The same uterus as in C dissected ex vivo; the tissue is viewed from the ventral surface. (E) Histology of the uterine mass (H&E stain), indicating adenocarcinoma from this example. For A and B, in terms of orientation, the mouse is dorsal surface up, and cranial to caudal is left to right (MS550 transducer). Scale bars = (A,B, first two images in each row) 1 mm; (A,B, third image) 2 mm; (E) 500 µm. Abbreviations: iPAD = inducible Pten, Arid1a double deletion; H&E = hematoxylin and eosin. Please click here to view a larger version of this figure.

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Discussion

This protocol examines the utility of ultrasound for assessing uterine morphological changes in the progression of adenocarcinoma in the uterus in mice. In this study, by following the induction of endometrial cancer in mice longitudinally, the anatomical details detected by ultrasound were found to be indicators of gross and histological pathology. This opens the door for the use of longitudinal studies with smaller numbers of mice monitored by ultrasound at multiple time points to follow the progression of uterine cancers in mice. This longitudinal detection was accomplished by using a hand-held probe and without the use of rail system ultrasound equipment. The high-frequency probes (transducers) used are widely available, making this approach reproducible at multiple institutions worldwide. Critical steps in the protocol include using the correct probe for the level of resolution required for the experiment and preparing the mouse for imaging prior to the ultrasound.

Critical steps in preparing the mouse are removing all the hair from the study areas and injecting fluid into the peritoneal cavity. Fur left on the skin can lead to the trapping of bubbles in the ultrasound gel, which can produce artifacts in the images; hence, care should be taken to remove the hair evenly over the study area. Depilatory cream was chosen as the method for hair removal as it removes the fur evenly across the skin. Injury to the skin can occur if the cream is left on the skin for an excessive amount of time (>3-4 min). In mice more sensitive to depilatory cream, the cream should only be left on the skin for 1-2 min to remove the hair. Next, the mouse should be wiped with a series of wet paper towels to completely remove the cream. After all the hair has been removed from the study area, sterile, warm isotonic fluids are injected into the intraperitoneal cavity to increase the contrast and assist in defining the boundaries of the abdominal organs. In this study, injecting 1-2 mL of fluid was sufficient to differentiate the ovary, oviduct, and uterine horns. The fluid is gradually absorbed, and it is best to inject immediately before imaging. Mice become hypothermic rapidly under isoflurane anesthesia. This can be prevented by injecting warm saline (stored in syringes on a heating pad) and by also positioning the mouse on a heating pad and prewarming the ultrasound gel when possible. Imaging sessions with experienced sonographers run at 5-10 min/mouse, and with this short time, significant hypothermia did not occur. If required, up to an additional 2 mL of saline can be injected into the abdomen for greater separation of the organs.

The methods described here are intended to be readily reproducible without specialized equipment beyond the ultrasound machine and probe(s). In the presented method, the probe is manipulated manually throughout without assistance from a rail system. This manual method is faster and provides great flexibility in the angles and pressure applied to highlight small features. Manually directing the probes over the abdomen and the use of lower-resolution transducers could limit the quality of any individual images acquired as compared with images from a transducer fixed on the rail system4. A rail system would be useful for injections into the uterus (into fetuses) or intracardiac injections when more stability is needed. However, there is a trade-off regarding minimal manipulation time and increased accessibility for the wide range of institutions that lack specialized ultrasound facilities. The success of this simplified method exemplifies how ultrasound can be utilized for a range of studies. The proposed method is well suited for capturing longitudinal tissue changes in uterine cancer models without the requirement for dedicated equipment and could be applied to other progressive disease models. Manual manipulation of the mice during imaging can increase the variability in the exact tissue depth and location, but for studies not relying on these factors, the image quality is sufficient to detect pathological changes.

It is important to save videos during imaging; later at the user's workstation, the videos can be stopped at any frame that is representative of the anatomy that needs to be captured and measured. Images of the organs can be measured (diameter, circumference, or length) (Figure 5, uterine horn cross-section measured), and the tumor area can be calculated. Videos, more so than single frames, facilitate the identification of organs in greater anatomical context. Videos are also useful for determining the movement of the small and large intestines, which can help differentiate the intestines from the uterus. Obtaining high-quality images also depends on familiarity with mouse anatomy. For imaging the movable uterus versus imaging more stationary tissues (e.g., an implanted xenograft tumor, ovaries), using the manual method has the advantage of scanning and following the uterine horns from proximal to distal. During scanning, gradual adjustment of the hand angles can be used to follow the horn. The MS550 probe was used for all of the imaging in this study and is preferable for the uterus compared to the higher-resolution MS700 probe, which is better for ovaries. Work using the MS700 to image the mouse thyroid has recently been published, showcasing the high resolution that is possible in other tissues15. Hand-held probe imaging is helpful due to the varying degrees of tissue depth of the uterus between the ovary and the bladder. In this work, moving between easily recognizable anatomical landmarks, such as the kidney and the proximal hind-limb anatomy, aided in capturing comparable images across the different time points when imaging the uterus from the dorsal aspect on the body.

This high-resolution ultrasound method to follow the anatomical changes in the uterus over time during tumorigenesis is superior to the current methods that involve euthanizing animals from multiple cohorts at different time points. Ultrasound refines the model and reduces the number of mice needed for the experiment, thus reducing costs and time. Importantly, this method allows for disease progression to be followed on an individual level, which may lead to insights that are masked in studies dependent on cohorts sacrificed by time point rather than disease stage. Due to its non-invasive nature, mouse ultrasound has many potential applications for research in uterine or ovarian cancer.

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Disclosures

The authors have no conflicts of interest to disclose.

Acknowledgments

We are grateful for funding from the NCI Ovarian Cancer SPORE Program P50CA228991, post-doctoral training program 5T32OD011089, and the Richard W. TeLinde Endowment, Johns Hopkins University. The project was also partly funded by the subsidies for current expenditures to Private Institutions of Higher Education from the Promotion and Mutual Aid Corporation for Private Schools of Japan.

Materials

Name Company Catalog Number Comments
Reagents and Equipment Used for Animal Care
Rodent Diet (2018, 625 Doxycycline) Envigio TD.01306 Mouse Feed
Reagents and Equipment Used for Ultrasound Imaging
10 mL injectable 0.9% NaCl  Hospira, Inc RL-7302 Isotonic Fluid
Absorbent Pad with Plastic Backing Daigger EF8313 Absorbant Pads
Anesthesia Induction Chambers Harvard Apparatus 75-2029 Induction Chamber
Anesthetic absorber kit with absorber canister, holder, tubing, & adapters CWE, Inc 13-20000 Nose Cone and Tubing
Aquasonic Clear Ultrasound Gel (0.25 Liter) Parker Laboratoies 08-03 Ultrasound Gel
BD Plastipak 3 mL Syringe BD Biosciences 309657 Syringe
F/Air Scavenger Charcoal Canister OMNICON 80120 Scavenging System for Anesthesia
Isoflurane, USP Vet One 502017 Anesthesia Agent
M1050 Non-Rebreathing Mobile Anesthesia Machine Scivena Scientific M1050 Anestheic Vaporizer
MX550S, 25-55 MHz Transducer, 15mm, Linear VisualSonics MX550S Ultrasound Transducer (Probe)
Nair Hair Aloe & Lanolin Hair Removal Lotion - 9.0 oz Nair Depilliating Cream
Philips Norelco Multigroomer All-in-One Trimmer Series 7000 Philips North America MG7750 Clippers
PrecisionGlide 25 G 1" Needle BD Biosciences 305125 Needle
Puralube Ophthalmic Ointment Dechra 17033-211-38 Lubricating Eye Drops
Vevo 3100 Imaging System VisualSonics Vevo 3100 Ultrasound Machine
Vevo LAB 5.6.1 VisualSonics Vevo LAB 5.6.1 Ultrasound Analysis Software
Vinyl Heating Pad with cover, 12 x 15" Sunbeam 731-500-000R Heating Pad
Wd Elements 2TB Basic Storage Western Digital Elements WDBU6Y0020BBK-WESN Data Storage
Reagents and Equipment Used for Immunohistochemistry
10% w/v Formalin Fischer Scientific SF98-4 Tissue Fixation Buffer
Animal-Free Blocker and Diluent, R.T.U. Vector Laboratories Inc.  SP5035 Antibody Blocker
Charged Super Frost Plus Glass Slides VWR 4831-703 Tissue Mounting Slides
Citrate Buffer MilliporeSigma  C9999-1000ML Epitope Retrival Buffer (pTEN)
Cytoseal – 60 Thermo Scientific 8310-4 Resin for Slide Sealing
Gold Seal Cover Glass Thermo Scientific 3322 Coverslide
Harris Modified Hematoxylin MilliporeSigma HHS32-1L Counterstain Buffer
Hybridization Incubator (Dual Chamber) Fischer Scientific 13-247-30Q Oven to Melt Parraffin
ImmPACT DAB Substrate, Peroxidase (HRP) Vector Laboratories Inc. SK-4105 Signal Development Substrate
ImmPRESS HRP Goat Anti-Rabbit IgG Polymer Detection Kit, Peroxidase Vector Laboratories Inc. MP-7451 Secondary IHC Antibody
Oster 5712 Digital Food Steamer Oster 5712 Vegetable Steamer for Epitope Retrival
rabbit mAB anti-ARID1a abcam ab182560 Primary IHC Antibody (1:1,000)
rabbit mAB anti-PTEN Cell Signaling 9559 Primary IHC Antibody (1:100)
Scotts Tap Water Substitute MilliporeSigma S5134-100ML "Blueing" Buffer
Tissue Path IV Cassette Fischer Scientific 22272416 Tissue Fixation Cassette
Trilogy Buffer Cell Marque  920P-10 Epitope Retrival Buffer (ARID1a)

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Vistein, R., Winer, B., Myers, S., Liberto, J., Ishiyama, S., Guo, X., Saeki, H., Wang, T. L., Shih, I. M., Gabrielson, K. Non-Invasive Ultrasound Assessment of Endometrial Cancer Progression in Pax8-Directed Deletion of the Tumor Suppressors Arid1a and Pten in Mice. J. Vis. Exp. (192), e64732, doi:10.3791/64732 (2023).More

Vistein, R., Winer, B., Myers, S., Liberto, J., Ishiyama, S., Guo, X., Saeki, H., Wang, T. L., Shih, I. M., Gabrielson, K. Non-Invasive Ultrasound Assessment of Endometrial Cancer Progression in Pax8-Directed Deletion of the Tumor Suppressors Arid1a and Pten in Mice. J. Vis. Exp. (192), e64732, doi:10.3791/64732 (2023).

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