Breast cancer is the most prevalent cancer and the second leading cause of cancer-related death for women in the US.Current prevention approaches are limited and come with a variety of adverse side effects. New interventions for primary prevention are needed. We have developed a method for intraductal delivery of an ethanol-based ablative solution to the mouse mammary epithelia for breast cancer prevention.
Injection directly into the nipple opening allows for targeting epithelial cells lining the ductal tree with minimal collateral tissue damage. This method allows for simultaneous in vivo imaging using tantalum oxide nanoparticles as a high-resolution contrast agent. This method may be adapted for use with other x-ray contrast agents and various preventative treatments with different chemical or thermal ablative solutions.
The first step involved with this method of image-guided breast cancer prevention is to induce extended anti-inflammatory treatment through oral dosing of carprofen utilizing sucralose gel cups prepared in the lab. We prepare the animal by using depilatory cream to remove the fur from the area surrounding the nipples to be injected. Then intraductal injections are performed with the help of a stereoscope to best locate and visualize the opening of the nipple for needle insertion.
After delivery of the solution, animals are imaged by micro-CT or fluoroscopy to determine how successful the injections were. Micro-CT scans can be processed to create 3D reconstructions of the injected mammary ductal tree for further analysis of successful filling and other metrics. First, prepare carprofen at required concentration.
We make a working solution of two mg per mL to inject 0.5 mL and achieve a final dose of one mg per cup. You may add sterile food dye to the mixture to better confirm full mixing of the drug into the sucralose of the cup. Next, place the cups into a water bath at 60 degrees Celsius for 15 minutes.
Remove the cups and dry them off to minimize possibility of contamination. Use 70%ethanol or ethanol wipes to clean the surface of the cup lid and allow to dry. Use a syringe to inject necessary amount of carprofen solution through the lid.
In this case, we need to inject 500 microliters for the desired result. Place a sticker over the hole from the needle and shake the cup for 15 seconds before vortexing for an additional 15 seconds. Check the cup for homogenous mixing by looking for clumps of dark blue.
Cups may then be refrigerated for up to a month. Preparation of the animal should occur two to three days before intraductal procedure. Begin by anesthetizing the animal using isoflurane.
Transfer anesthetized mouse to a nose cone which continues to deliver isoflurane. Apply eye lubricant to the mouse while at the nose cone and then position the mouse on its back. Use a cotton tipped applicator to apply depilatory cream to the area of the mammary glands you plan to inject.
Rub the applicator up and down across the animal's skin in the desired area to help with quicker loosening of the fur. It is important to leave the cream on the mouse for as little time as possible to avoid burning the skin. After 10 to 30 seconds of application, use warm water on gauze to completely remove the cream.
Use two to four rinses to ensure complete removal before drying the skin with clean gauze. Check the areas of fur removal to confirm good visibility and access to the nipples. Repeat depilatory procedure if necessary.
Place the mouse in a clean recovery cage on a warming pad to recover from anesthesia. Give one carprofen cup to prep mice in their home cage after recovery. You will again begin by anesthetizing the animal using isoflurane anesthesia and move the mouse to a nose cone once it has been fully induced.
Eye lubricant will be applied before placing the animal on its back for injections. It can be helpful to tape down the legs near the mammary glands that will be injected, but is not necessary. Prepare the syringe with the desired volume of injection solution plus one microliter in case some spills out after needle removal.
It can be helpful to fill with additional volume to test the needle for free flow of the solution immediately prior to injection. If injecting more than one mammary gland, it can save time to prefill multiple syringes if available. Take care not to do this too early if the solution is prone to clogging when left to sit.
Prepare the nipple by removing any visible dead skin using fine tipped forceps if possible. It is not necessary to remove all dead skin if it is not blocking the nipple opening and it is resistant to tugging. This can sometimes cause injury and swelling to the nipple if dead skin is affixed to live skin too tightly.
With the bevel of the needle visible, insert the needle into the opening of the nipple with guidance help from the fine tipped forceps. Depending on the length and firmness of the nipple, it may not always be possible to hold the nipple while guiding in the needle. It is sometimes necessary to maneuver the nipple with the needle to allow for the forceps to pull the nipple up over the needle rather than guiding the needle down into the nipple.
Once the needle bevel is fully surrounded by the nipple and in the main duct, begin slowly injecting the solution. The desired rate is approximately 40 microliters per minute. Avoid injecting more quickly than this to prevent possible damage to the ductal tree.
After complete injection, wait 30 seconds before removing the needle from the nipple with assistance from the forceps. This will minimize any leakage from the nipple. Assess the area for any signs of an unsuccessful injection.
A domed appearance may indicate a fat pad injection or trauma to the area. It is often possible to see that an injection into the fat pad is occurring if the solution contains dye or if you are looking carefully for doming at the injection site as can be seen here. Proceed to inject the remaining mammary glands required for your experiment in similar fashion, noting injection volumes and any incidences of leakage.
Some areas of the mouse may be more difficult to inject due to issues such as awkward hand placement or breathing interference. For example, a right-handed injector will likely have the hardest time injecting the glands on the mouse's upper right side seen here. These move more due to proximity to the lungs and it can be difficult to find good hand placement with the body of the animal in the way of resting the wrist on the surface.
Some may find it easier to reposition the mouse if the nose cone allows. When injecting an ethanol-containing solution, care must be taken to avoid systemic alcohol intoxication. This requires knowing how much ethanol can be injected in one session and administering sucrose-containing solution IP during the procedure to counteract the effects.
After intraductal procedure, animals may be recovered in a clean cage on a heating pad or moved to the micro-CT system for imaging of filled ductal trees. The mouse should also be recovered following any imaging session. After moving the injected animal to the micro-CT system, continue administering isoflurane to maintain anesthesia during imaging.
It can be helpful to standardize imaging position by taping down the legs near the site to be imaged. For example, taping the hind legs down in an extended position can keep the bone of the legs from impeding the view of the injected glands. Taping the front legs in an upward extended position can similarly aid in viewing the upper glands.
We have also found that taping across the abdomen of the mouse can assist in reduction of breathing artifact for the lower glands. There are many acceptable scanning parameters for visualization of the ductal tree. Care should always be taken to ensure the radiation dose resulting from these scans does not exceed radiation limits for a given strain of mice.
Radiation dosing can be greatly reduced by viewing fluoroscopy and noting delivery success rather than acquiring images with longer exposure times. Our standard scan parameters are acceptable for repeat imaging if only acquiring short two-minute standard scans. Higher resolution longer scans up to 14 minutes are acceptable as terminal procedure before euthanasia.
These longer scans are not acceptable for longitudinal studies as they can cause radiation sickness in the animal. This animal was injected with an ethanol solution containing 100 millimolar tantalum oxide contrast in glands four and nine. The current acquisition is a 14-minute high-resolution scan of the injected lower glands.
Fluoroscopy of the animal allows us to see the ductal tree architecture being highlighted by the contrast within the injection solution. The micro-CT system we use has built-in software that allows for creation of quick renditions to assess the success of injection without formal analysis. This feature has a simple contrast slider that allows for reasonable signal-to-noise reduction.
The limitation of these renditions is that the same signal thresholds are applied to the entire image. This allows bright signals that are clearly not part of the ductal tree, such as iron in the diet, to remain in the rendition. Better formal renditions can be made using more sophisticated analysis software that allows segmentation of the area of interest.
To get the best rendering of the sole ductal tree, we recommend segmenting out the mammary fat pad for further image processing. This process requires line tracing the boundaries of the fat pad throughout the complete thickness of the animal. We have found that tracing every third slice and propagating the object is sufficient to capture the entirety of the duct.
This can be a subjective process, which requires adherence to lab standards for tracing to ensure similar output from all individuals analyzing scans. Familiarity with anatomy can help with judgment calls about what areas to include. After propagating traces, it is then possible to threshold the rendition within a certain range to display only the contrast contained within the mammary gland.
This should be the solution within the ductal tree, as well as any leakage in the immediate vicinity. A range of 300 to 3, 000 HU tends to work well in the case of tantalum oxide-containing solutions. An example of a fully processed rendition can be seen here in multiple aspects.
Depending on how the animal is oriented in the micro-CT, it may be necessary to reorient DICOM files in the analysis software to represent intuitive anatomical positioning with the head of the animal at the top of the image. It may also be difficult to determine which gland is being viewed if you do not take care to rotate the image in similar fashion each time. Further measurements such as volume or length can be made once you have created a reconstruction of the ductal tree.
Validation for successful delivery through this procedure can be demonstrated not only by 3D micro-CT renditions as described here, but also through dual-stain whole mounts with injected Evans blue overlapping the entire ductal tree highlighted with carmine alum. 3D confocal microscopy can highlight the architecture of the ductal tree in a cleared gland stained with e-cadherin. Strain-to-strain and mouse-to-mouse variability and nipple anatomy can make it more or less difficult to inject glands.
This is an example of an ideal nipple for injection with a high profile for ease of gripping. Nipples with lower profiles, such as this one, can be more difficult to cannulate successfully. These differences can make it more likely to achieve a successful injection with no trauma or doming as seen here or more likely to be unsuccessful as shown by the domed appearance of this fat pad injection.
When it comes to breast cancer prevention, there are currently two main tactics being employed. The first, watchful waiting, involves no true intervention. Individuals pursuing this path are regularly monitored for any lumps in their breast tissue that would indicate cancer formation is occurring.
Disease detection would then prompt treatment. Prophylactic mastectomy is a more invasive procedure which removes all epithelial cells from which breast cancer may arise along with the surrounding stroma. This surgery can have serious adverse side effects and is rarely selected as a prophylactic option for individuals with low or moderate risk of disease.
Our approach involves local epithelial ablation through injection of an ablative solution directly into the mammary ductal tree. This method reduces collateral damage and would likely be a more appealing option for the individuals not at severe risk of disease. The addition of contrast for immediate imaging post-injection also makes this easily translatable to the clinic.
