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

Summary

This protocol describes a method for monitoring the progression of morphological changes over time in the uterus in an inducible mouse model of endometrial cancer using ultrasound imaging with correlation to gross and histological changes.

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 disorders^1,2,3. 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 animal^4,5. 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 resolution^6,7. 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 neoplasia^9,10,11. 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.

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. Apply sterile ophthalmic lubricant to each eye. 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. Apply sterile ophthalmic lubricant, as needed, to each eye.
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.

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 c...

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 canc...

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)