Sectioning Mammary Gland Whole Mounts for Lesion Identification

We developed a method to successfully remove, process, section, and stain, for histopathological evaluation, mammary tissue that had originally been fixed on slides as whole mounts. This method may promote the collection and evaluation of mammary gland whole mounts in reproductive and developmental test guideline studies.

Transcript

The overall goal of this procedure is to create high-quality stained sections from whole mounted mouse mammary tissue for lesion identification at the microscopic level. This method can help answer key questions involving mammary gland evaluation such as histopathological identification of lesion types identified grossly in mammary whole mounds. The main advantage of this technique is that it increases the chance of identifying lesions that may exist within the mammary glands when using whole mounted tissues.

We first had the idea for this method when we compared whole mount phenotypes to histopathology reports for formalin fits H and E stain contralateral glands from the same animals. We saw a discrepancy in diagnoses, more lesions were evident in whole mounts. Visual demonstration of this method is important as the whole mount removal and embedding steps are critical to the success of this procedure.

The gland is delicate because of its thin friable nature. After preparing whole mount memory samples and analyzing the tissue for abnormalities according to the text protocol, remove the cover slips and mounting medium by submerging the slides in a glass staining jar filled with xylene. Incubate overnight or longer.

Transfer the slides to fresh xylene and soak the samples for six hours. Then transfer the samples again to fresh xylene and soak overnight. Next, with a gloved hand, hold the slide and use a pair of forceps to carefully remove the coverslip in one piece.

There may be variability in the ease of removing the coverslip depending on the thickness of mounting medium and the time since the whole mount was coverslipped. Therefore it is imperative that the slide remain in xylene until the coverslip is easily released. Once the coverslip is removed, use a sharp disposable razor blade to remove the sample by sliding the blade in a single motion down the slide.

Then quickly submerge the memory pad into a xylene filled glass petri dish to ensure that the tissue does not air dry. Use a razor to cut larger tissues in half. Then to minimize tissue damage, use blunt nosed serrated-tipped forceps to grab the edge of the fat pad and place the halves right side up in two separate labeled cassettes that will accommodate the tissue size.

Place the cassette into a container of xylene for up to two hours before placing it in the tissue processor. Then load the maximum number of cassettes into the tissue processor by hand. Prior to beginning, program the processor for tissue processing.

To automate the procedure, select the station, the temperature and vacuum on off on. Automate the processor to soak the cassettes in xylene for 30 minutes at 37 degrees celsius followed by a second soak in fresh xylene under the same conditions. Then automate the processor to remove the cassettes from the xylene solution and transfer the tissues to a chamber within the processor containing a one-to-one xylene to molten paraffin mixture set at 60 degrees celsius for 30 minutes.

When all the steps have been completed, select okay or start to proceed. Place the cassettes in the holding tank of a 58 degrees celsius paraffin bath until ready to place them in the mold. Remove cassette covers to determine the best mold size for the tissues.

Add three to four millimeters of molten paraffin from the spout to the mold, then orient the tissue in the molten paraffin so that the mammary whole mount surface which was adjacent to the bottom of the glass slide and the bottom of the cassette, is facing up in the mold. Transfer the mold to a cooling plate and quickly adjust the tissue as needed so that it is parallel to the mold bottom. Using forceps, place the labeled cassette firmly on top of the mold.

Dispense additional molten paraffin into the mold in a continuous motion to cover the entire cassette. Move the mold from the warm plate to a cold plate to promote fixation in the exact position chosen for sectioning. When the paraffin block completely solidifies, separate the block from the mold.

And store it at room temperature until ready for sectioning. Prior to sectioning, incubate the paraffin blocks at minus 20 degrees celsius for one hour. Then prepare the microtome by turning on the water bath containing fresh distilled water and adjust the temperature to 42 to 45 degrees celsius.

Next, place a fresh low-profile blade onto the microtome and set it at four micrometers. Insert the block into the microtome with the wax facing the blade and aligned with the vertical plane. Then moisten a section of gauze pad in cold water and place it onto the block for several minutes.

Section the block by turning the large wheel in a clockwise motion in combination with the course advanced wheel until a full face or representative section of block is obtained. Then transfer the tissue section by hand to a warm water bath before using a clean slide to pick it up. Place the slides on a drying bracket to remove excess water.

Then store the tissue sections at 37 degrees celsius overnight. After removing the slides from the 37 degree celsius incubator, store them at room temperature until ready for staining. For automated staining, select H and E from the stain options on the main menu to stain the slides.

Deparaffinization, staining and dehydration are all completed on the automated stainer. Use mounting medium and a coverslip to mount the sections. To determine the ideal thickness of the mammary tissue four micrometer and six micrometer sections were prepared.

The four micrometers sections produced optimal results where epithelial and stromal areas and corresponding cell types were easily distinguished. These figures represent histological sections of normal glands. Both glands show ductal structures surrounded by a robust and homogenous adipocyte-rich population.

Each duct is lined by a single layer of simple cuboidal epithelial cells and is maintained by a second layer of basal cells. The contralateral whole mounts contained an area of increased opacity that involved the memory glands ducts. However, it was unclear if the opaqueness was from a hyperplastic, inflammatory or neoplastic alteration.

Contralateral whole mounts are shown here. These sections were diagnosed as perivascular inflammation due to the increased number of lymphocytes around a large blood vessel in the mammary glands section. In these samples, the mammary gland lobular architecture was maintained, but was enlarged by increased numbers and size of normal alveoli and ducts for lobular alveolar hyperplasia.

Enlarged lobules contained an increased number of alveoli and ducts that were lined by well-differentiated, often vacuolated epithelial cells that formed lumens that contained proteinaceous fluid. While attempting this procedure, it's important to remember that the tissue is fragile from the xylene and it may easily break, destroying your sample. High-quality photos of whole mounts using a flatbed scanner are encourage before attempting this method.

This is an opportunity for the field of toxicological pathology to move in the direction of whole mount evaluation for chemical screening. So when lesions are identified at that level, high-quality sections can now be produced for histopathologic diagnosis. After watching this video, you should have a good understanding of how to remove, embed and section mammary whole mounts for lesion identification.

Don't forget that when working with xylene, precautions, such as lab coats and gloves should be worn and the work should take place under a hood.

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Journal (Cancer Research)

Journal (Cancer Research)

Sectioning Mammary Gland Whole Mounts for Lesion Identification

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