X-Ray Visualization of Intraductal Ethanol-Based Ablative Treatment for Prevention of Breast Cancer in Rat Models

Breast cancer is a prevalent and deadly disease with limited prevention strategies that come with severe negative side effects. To provide a method with reduced side effects, we have developed a technique for intraductal injection of an ethanol-based ablative solution to the rat mammary ductal tree that allows for simultaneous in vivo imaging and breast cancer prevention. This builds on previous work in mice in which injection directly into the nipple opening allows for targeting mammary epithelial cells with minimal collateral tissue damage.

Both mice and rats have mammary glands containing a single ductal tree originating from the nipple opening. Rats are larger than mice and allow for injection of a larger volume of ethanol for study of ablation success to prove scalability of the model. Another refinement to this method is the addition of ethyl cellulose to the ablative solution.

Ethyl cellulose has been used previously in other clinical approaches to reduce the spread of ethanol away from the area being targeted. This is accomplished through the nature of the compound, which causes it to gel when it contacts fluid in the tissue. There are five key steps involved in this method of image verified breast cancer prevention.

We must first begin dosing animals with carprofen orally utilizing sucralose gel cups prepared in the lab. This extended anti-inflammatory treatment is maintained pre and post procedurally. Next, we remove the fur from the area surrounding the nipples to be injected with depilatory cream to prepare the animal for injections.

Intraductal injections are then performed with a stereoscope aiding visualization of the nipple opening for needle insertion. Animals are scanned or assessed by microCT or fluoroscopy following injection to determine success of injections. microCT scans may then be processed to create 3D reconstructions of the injected mammary ductal tree for further analysis of successful filling.

Stock carprofen is first diluted in PVS to the required concentration. Sterile food dye may be added to the mixture to better confirm full mixing of the drug into the sucralose of the cup. Cups are then heated in a water bath at 60 degrees Celsius for 15 minutes.

Drying the cups upon removal and cleaning the cup lid with 70%ethanol reduces risk of contamination. Carprofen solution can be injected directly into the cup using a syringe to pierce the lid. In this case, we need to inject 500 microliters for the desired result.

Seal the hole with a sticker before shaking the cup for 15 seconds and then vortexing for an additional 15 seconds. Homogenous mixing can be assessed by looking for clumps of dark blue. Cups can then be refrigerated until needed.

Animals should be prepared two to three days before injections are to occur. Anesthetize the animal using inhalant isoflurane. Transfer anesthetized rat to a nose cone on a warming pad.

Apply eye lubricant to the rat while at the nose cone and then position the animal on its back. Depilatory cream can be applied to the area of the nipples you plan to inject with a cotton tipped applicator. Quicker loosening of the fur can be achieved by rubbing the applicator up and down across the animal's skin in the desired area.

It is important to leave the cream on the rat for as little time as possible to avoid burning the skin. Rats are even more sensitive than mice to this procedure. After 10 to 30 seconds of application, use warm water on gauze to completely remove the cream.

Use three to four rinses to ensure complete removal before drying the skin with clean gauze. Confirm good visibility and access to the nipples in the areas of fur removal. Repeat depilatory procedure if necessary.

Place the rat in a clean recovery cage on a warming pad to recover from anesthesia. Give one carprofen cup to prepped rats in their home cage after recovery. You will begin by anesthetizing the animal using inhalant isoflurane anesthesia and move the rat to a nose cone once it has been fully induced.

Eye lubricant should be applied before placing the animal on its back for injections. It can be helpful to tape down the legs near the nipples that will be injected but is not necessary. Once you have prepared the syringe with the desired volume of injection solution, prepare the nipple by removing any visible dead skin using fine-tipped forceps if possible.

The most common area of concern is the nipple opening itself. Rats often have a plug protruding from the opening. Removal of this plug can aid in better cannulation.

With the bevel of the needle visible, insert the needle into the tip of the nipple with guidance help from fine-tipped forceps. Take care to follow the path of the nipple opening. Rat nipples tend to have more fat surrounding the opening making it easier to pierce into this fat and mistakenly believe you have achieved cannulation due to advancement of the needle.

Once the needle bevel is fully surrounded by the nipple begin slowly injecting the solution. The desired rate is approximately 100 microliters per minute in rats. 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 lessen the likelihood of leakage from the nipple. If leakage does occur, use moistened gauze or an ethanol wipe to clean off the solution.

Contrast and spilled solution may distort acquired images. Here we see a magnified video of the injection sequence that was just displayed. Notice that the animal's breathing makes it difficult to keep the nipple in a single focal plane.

This can make injection more difficult as the nipple does not remain in a constant location. This problem is exacerbated by proximity to the ribcage and the top three gland pairs. The plug in the nipple opening has now been removed for better cannulation.

The needle may now be oriented bevel up to be advanced into the nipple with the assistance of the forceps. Once the bevel is fully inserted, injection can begin. If you look carefully to the left side of the nipple you may notice an increase in blue tint where the injection solution is slightly visible spreading through the ductal tree.

This is much less visible in the rat due to thicker skin. Here we can see removal of the needle 30 seconds after injection has completed and the resultant solution spilling out of the nipple. The extra solution is cleaned off, allowing assessment of the area where we see no trauma to the nipple or doming of the area, which would indicate a fat pad injection.

If injecting a solution containing ethanol, care must be taken to avoid 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. Following injection, animals may be recovered in a clean cage on a heating pad or moved to the microCT for imaging of injection success.

After moving the injected animal to the microCT instrument, continue administering isoflurane to maintain anesthesia during imaging. The increased size of the rat requires additional position manipulation to achieve a good image. Straightening of the spine before taping down the legs near the site to be imaged is helpful in generating consistent images.

Taping legs in an extended position can prevent bones from becoming the focal point of the image. We have also found that taping across the abdomen of the rat can assist in reduction of breathing artifact for the lower glands. Several scanning parameters are acceptable for visualization of the ductal treat.

However, radiation dose should always be considered when selecting parameters. Radiation dose resulting from these scans should not exceed radiation limits for a given strain of rats. Fluoroscopic analysis, rather than image acquisition, can greatly reduce overall radiation burden.

Once all scans or assessments have been made, the animal may be returned to a clean recovery cage on a warming pad. The animal should be monitored until fully recovered and returned to the home cage. Carprofen cups should be provided until at least seven days post-injection.

The software on the microCT instrument we use 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 primary flaw in these renditions is that the entire image must be thresholded simultaneously.

This allows bright signals that are clearly not part of the ductal tree, such as iron in the diet, to remain in the image. In some cases, the true signal as seen here is as dim as background and can easily be thresholded out of the image. Better formal renditions can be made using more sophisticated analysis software that allows segmentation of the area of interest.

It is best to segment out the mammary fat pad for further image processing in order to get the best rendition of the injected ductal tree. The dark boundary of the mammary fat pad may be spline traced throughout the complete thickness of the animal in order to achieve this segmentation. Tracing every third slice and propagating the object is sufficient to capture the entirety of the duct in our experience.

At this point, it is possible to threshold the rendition using a given rage to display only the contrast contained within the mammary fat pad. This should include the solution within the ductal tree as well as any leakage in the immediate vicinity. Tantalum oxide solutions are generally well displayed in a range of 300 to 3000 HU.Further analysis can be formed once you've created a reconstruction of the ductal tree.

If longitudinal imaging is not desired, higher resolution images that better capture the complex architecture of the ductal tree may be acquired. Differences between animals can make injection more or less difficult. We cannot speak to strain to strain variability as we currently exclusively work with Sprague Dawley rats.

Some animals will have nipples with low profiles, like the one pictured here, making it difficult to manipulate and achieve successful cannulation. Others will have nipples with a higher profile, like the one shown here, that will be more amenable to cannulation. A successfully cannulated nipple is depicted here.

Difficulty cannulating can result in trauma to the nipple as shown here. Acquisition of microCT images immediately post-injection can better inform researchers of the success of an injection as well as help to identify filling properties of certain solutions. Here we see different aspects of abdominal glands of the same rat injected with the same volume of an ablative solution containing 70%ethanol with contrast provided by 100 millimolar tantalum oxide.

The solution used in the top gland shown also contained 1%ethyl cellulose. We see wall-like structures at the ends of the ductal branches. These are terminal end buds filled with the solution and not spilling over.

In the bottom row, we see less defined ductal ends indicating the escape of the solution from the ductal tree. This, along with more limited collateral damage in histological examination. seems to indicate better retention of solutions containing ethyl cellulose to the ductal tree.

This method provides a step towards scalability of a less invasive method of breast cancer prevention to offer an alternative for prophylactic mastectomy. Successful epithelial ablation in a larger rodent model has been demonstrated with minimal side effects to the animal. Addition of ethyl cellulose to the ablative solution allows for better retention of the solution to the area of injection, which results in reduced collateral damage.

The novel use of solutions that have been clinically proven and added ability to visualize delivery success with common imaging modalities allows for ready translation into the clinic.