Isolation of Intact, Whole Mouse Mammary Glands for Analysis of Extracellular Matrix Expression and Gland Morphology

The overall goal of this whole mammary gland harvest technique is to facilitate the analysis of the histological presentation of mammary tissue from an intact gland ex-vivo. This method can help answer key questions in the tumor biology field, such as how stromal tissue is remodeled in tumors or how aberrant gene or protein expression alter mammary ductal architecture. The main advantage of this technique is that it allows researchers the ability to characterize and quantify protein expression and ductal architecture in whole mammary glands of healthy and diseased mice.

Demonstrating this technique will be Chris Thompson, a graduate student in my laboratory. After sacrificing eight to 10 week old female nulliparous mice according to the text protocol, pin the carcass in a supine position and use 70%ethanol to saturate it. Use forceps to pinch the pelt just above the pubis, and nick it with small surgical scissors.

Then, rotating the scissors, cut the pelt along the ventral mid-line, moving caudal to cranial. With larger scissors or a hemostat, bluntly dissect the subcutaneous fascia bilaterally, using caution not to puncture the peritoneum. Cut along the horizontal margins at both distal ends of the incision.

Next, pin the pelt flaps open and spray the tissue with PBS to keep it moist. Then, locating the mammary glands of interest, slide a scalpel blade along the inside of the pelt flaps, cutting the mammary gland and associated subcutaneous adipose free from the dermis. Before separating the abdominal gland from the inguinal, take care to identify the glands as distinctly as possible.

Immediately submerge the newly isolated gland in a conical tube containing approximately six milliliters of a 10 to one solution to tissue volume 10%neutral buffered formalin, and incubate it for 24 to 48 hours. After melting the paraffin off the slides and re-hydrating the slices according to the text protocol, boil 10 millimolar sodium citrate solution on a hot plate or in a microwave and pour the solution into a Coplin jar. Slowly place the slides into the Coplin jar, ensuring complete submersion, and incubate the slides at 100 degrees Celsius for 10 minutes to retrieve epitopes.

Then, remove and carefully dry the slides. Next, draw a barrier around tissue with a hydrophobic marker. Once it has dried, rinse the slides with PBS for one to two seconds.

Then, add enough endogenous enzyme blocker to cover the tissue, and incubate the slides for 10 minutes. Submerge the slides in 1X PBS for 10 minutes. Then, add enough 10%donkey serum and PBS to cover the tissue, and incubate the samples at room temperature in a humidified chamber for one hour.

Decant excess blocker from the slides and add properly diluted primary antibody in 1%serum directly to the slides, ensuring that the tissue is evenly covered in solution. Incubate the slides at room temperature in a humidified chamber for 30 minutes. Carefully wash the slides by flowing about one milliliter of deionized water over the tissue, and then incubate the slides in one change of 1X PBS for five minutes.

Decant the excess PBS and add enough horseradish peroxidase conjugated secondary antibody, and incubate the samples in a humidified chamber for 30 minutes. Carefully wash the slides by flowing about one milliliter of deionized water over the tissue, and incubate the tissue in one change of 1X PBS for five minutes as before. After preparing DAB plus chromogen solution, decant the excess buffer from the slides and add two to three drops of chromogen stain directly to the tissue.

Incubate the samples in a humidified chamber for five minutes. Then, carefully wash the slides by flowing about one milliliter of deionized water over the tissue. Following a final rinse after chromogen incubation, submerge the slides in Harris hematoxylin for two to two and a half minutes.

Then, gently rinse the slides under tap water for one to two minutes. Submerge the slides in differentiation solution for two to three seconds. Then, gently rinse the slides under tap water for about one to two minutes.

Submerge the slides in blue agent for 60 seconds. Then, rinse the slides in 95%ethanol for 30 seconds. Transfer the slides into alcoholic eosin Y for two to three minutes before dehydrating the tissue in two changes of 95%ethanol for one minute each.

Then, finally, dehydrate the tissue once in 100%ethanol for one minute. Carefully dry the slides with a lint-free wipe. Then, apply one drop of synthetic, non-aqueous resin-based mounting medium to the slide and apply a cover slip.

To carry out ductal analysis, select slides stained for alpha SMA. Then, using a light microscope fitted with a camera-mounted objective, collect representative images at 20X magnification. To count ducts in each image, either count them manually or open ImageJ and select plugins.

Next, select analyze, and from the drop-down menu, choose the cell counter. Click initialize, and then highlight type one to begin labeling ducts. When finished, select results to view the ductal count.

Start by drawing a line over the image's scale bar with the line tool. Then, click analyze and set scale. In the next window, type the distance in known distance and the units in unit of length.

Then, click okay. Next, in set measurement options, check area and perimeter. Then, using the freehand polygon tool, draw a line around each duct at the myoepithelial compartment.

Select measure, and record the perimeter and area. Using the freehand polygon tool again, draw a line along the apical side of the ductal epithelium within the interior of the lumen. Then, select measure, and record the perimeter and area.

Subtract the area of the lumenal compartment from the area of the myoepithelial compartment to obtain the area of the ductal epithelium. Subtract the perimeter of the lumenal compartment from the perimeter of the myoepithelial compartment to obtain the circumference of the ductal epithelium. To carry out immunohistochemical analysis, upload bright-field images to an analytical software, such as ImageJ or Fiji Suite.

Using the plugin menu in ImageJ, open the IHC toolbox. In the select model combo box that opens, select the appropriate stain. Select the color option to isolate the stain.

This will open a result window. Use the color chooser slide to ensure proper isolation of the stain, without background inclusion or excessive stain exclusion. Go to image type, 16-bit, to convert the image to 16-bit.

Then, threshold the image by going to image, adjust threshold. Finally, go to analyze, set measurement to measure the resultant area and mean gray value, and choose analyze, measure, to collect the measurement. In these fixed sections of number four mammary glands, expression of collagen, fibronectin, and tenascin-C were analyzed.

The arrows indicate the presence of ducts in these tissues. To quantify morphological features of ducts, it is important to treat the histological sections with care, as improper handling may result in damaged tissues and ducts that are not measurable. An example of damaged tissue that cannot be used for analysis is shown here.

An intact tissue specimen where the ducts are measurable and quantifiable is shown in this panel. The ducts are not only characterized by the surrounding fibronectin-stained stroma, but also by the dense population of epithelial cells lining the lumen of the ducts. In this fibronectin stained section, several intact ducts contain branches indicated by arrows.

For the purpose of this analysis, these ductal branches should not be considered as separate ductal structures. While evaluating and measuring ducts, it is important to distinguish ducts from vascular structures. Vascular structures are indicated by red arrows, and ductal structures are indicated by black arrows.

Vascular structures can not only be identified by the presence of an endothelial monolayer lining the lumen, but also by the presence of anuclear, eosinophilic structures indicative of red blood cells within the lumen. Once mastered, tissue harvest can be done in approximately 15 minutes per animal, and section stains can be completed in about seven to eight hours, with slide images taken the next day if all is performed properly. While attempting these procedures, it is important to remember that the individual goals of your experiment may require optimizations at specific steps.

Following this procedure, other methods like second harmonic generation or coherent anti-Stokes Raman scattering can be performed in order to answer additional questions regarding tissue stromal composition or architecture. After its development, this technique paved the way for researchers in many fields, such as tumor biology, to explore tissue organization and protein expression patterns in numerous animal models of human disease and pathologies. After watching this video, you should have a good understanding of how to dissect and isolate mouse mammary glands.

You should also understand the procedures of immunohistochemical staining of tissue sections, including the use of counter-stains. Don't forget that working with animals and their tissues and chemicals such as xylenes and DAB chromogen can be extremely hazardous, and precautions such as appropriate husbandry training and the use of personal protective equipment should always be taken while performing these procedures.