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

Summary

Here, we present a protocol for the isolation of whole, intact mouse mammary glands to investigate extracellular matrix (ECM) expression and ductal morphology. Mouse #4 abdominal glands were extracted from 8-10 week old female nulliparous mice, fixed in neutral buffered formalin, sectioned and stained using immunohistochemistry for ECM proteins.

Abstract

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal architecture. Here, a small pocket below the skin was created using Mayo scissors, allowing separation of the glands within the subcutaneous tissue from the underlying peritoneum. Visualization of the glands was aided by the use of 3.5x-R surgical micro loupes. The pelt was inverted and pinned back allowing identification of the intact mammary fat pads. Each of the #4 abdominal glands was bluntly dissected by sliding the scalpel blade laterally between the subcutaneous layer and the glands. Immediately post-harvest, glands were placed in 10% neutral buffered formalin for subsequent tissue processing. Excision of the entire gland is advantageous because it primarily eliminates the risk of excluding important tissue-wide interactions between ductal epithelial cells and other microenvironmental cellular populations that could be missed in a partial biopsy. One drawback of the methodology is the use of serial sections from fixed tissues which limits analyses of ductal morphogenesis and protein expression to discrete locations within the gland. As such, changes in ductal architecture and protein expression in 3 dimensions (3D) is not readily obtainable. Overall, the technique is applicable to studies requiring whole intact murine mammary glands for downstream investigations such as developmental ductal morphogenesis or breast cancer.

Introduction

Breast cancer is characterized by a substantial degree of tissue fibrosis^1,2,3,4. Referred to as the ECM, this non-cellular entity is found in varying degrees in all tissues and is primarily comprised of a complex meshwork of fibrillar and non-fibrillar collagens, elastin, and glycoproteins in addition to various signaling molecules that are sequestered in this matrix. Under homeostatic conditions, the deposition and degradation of the ECM is tightly controlled.⁵ During breast tumorigenesis, the balance of ECM deposition and degradation is disrupted. As such, breast tumors have been reported to express abundant ECM proteins such as collagens, fibronectin and tenascin-C amongst others.⁶ The abnormal expression of these proteins in addition to increased patterns of matrix crosslinking has been documented to promote breast tumor progression, metastasis and therapy resistance^1,3,4,7,8,9.

To assess ECM composition and ductal morphology, isolation of intact mammary glands was performed. Here, we used female nulliparous mice deficient for caveolin-1, an integral membrane protein which has been linked to an aggressive breast tumor signature^10,11,12, and control female nulliparous B6 mice. Histological processing and staining of these tissues permitted the identification of several ECM proteins along with characterization of ductal morphology.

Overall, the isolation of whole, intact mammary glands gives researchers the opportunity to investigate tissue-wide morphological or cellular changes occurring in response to exogenous or endogenous factors. Drawbacks of the technique are associated with analyses of 2 dimension (2D) tissue sections as opposed to a 3D perspective, which would yield a more complete picture of the complex morphology of the ductal tree. Given the complexity of cell-cell and cell-ECM interactions that take place in the mammary gland, the isolation of whole, intact glands is advantageous for efficiently analyzing ductal morphology and protein expression in various regions of the murine mammary gland.

Protocol

Procedures involving animal subjects in this protocol were reviewed and approved by the Institutional Animal Care and Use Committee of the Philadelphia College of Osteopathic Medicine and all techniques were conducted under strict ethical guidelines.

1. Sample Procurement and Processing

1. Select appropriate animal subject and place into a CO₂ chamber. For this experiment, use 8-10 week old female nulliparous B6.Cg-Cav1tmMIs/J and C57BI/6J.
2. Turn on gas flow to 30-40%. Once the animal is visibly unconscious after about 2 min, open the gas valve to full pressure for an additional 5 min.
   NOTE: Confirm animal death by observing visible signs of breathing (e.g. movement of the chest) for a period of 10 min after cessation of CO₂ delivery. If the animal is still alive, it may be placed back in the CO₂ chamber. Cervical dislocation may also be used although it is not recommended as blood may accumulate around the mammary glands interfering with the dissection and results.
3. Following sacrifice, pin carcass in a supine position and saturate with 70% ethanol.
4. Pinch the pelt just above the pubis using forceps and nick with small surgical scissors. Rotating the scissors, cut the pelt along the ventral midline moving caudal to cranial.
5. 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.
6. Pin the pelt flaps open and spray again with 70% ethanol.
   NOTE: Although the use of 70% ethanol permitted better visual distinction between the gland and the surrounding subcutaneous tissue, be careful to avoid drying of the tissue as a result of use of this solution. To avoid drying, remove the gland in a time frame not to exceed 4 min. As an alternative to 70% ethanol, the investigator may also substitute a general isolate buffer, such as 1x PBS, to avoid tissue desiccation.
7. Locating the mammary glands of interest, slide a #4 scalpel blade along the inside of the pelt flaps, cutting the mammary gland and associated cutaneous adipose free from the dermis.
   NOTE: 3.5x surgical micro loupes may aid in easier visualization of the glands.
8. Immediately submerge newly isolated gland in conical tube containing 10:1 solution-tissue volume of 10% neutral buffered formalin for 24-48 h.
   NOTE: Depending on intent of study, many alternative fixatives may be used such as paraformaldehyde, ethanol for genomic studies, and commercially available RNA preserving buffers.
9. Follow institutional protocols for paraffin embedding, submit samples to a vendor for processing, embedding, and slicing/slide mounting. Section tissues at 5 µm.
   NOTE: Due to excess adipose surrounding gland, consider removing 100-200 µm of tissue before sectioning and staining.

2. Tissue Staining

1. Immunohistochemistry
   1. Place slides to be stained on heat block set at 58 ˚C for 1 h to melt paraffin.
      CAUTION: Melting paraffin wax may run off slide. Monitor closely or place wipe under slide to capture runoff wax.
      NOTE: This step is not essential and may be skipped. If results are not as expected, adding this step back may improve results.
   2. Incubate slides in a slide jar containing 100% xylene for 30 min ensuring complete submersion. Repeat once.
      NOTE: At this point, visual inspection should be performed to ensure that tissue sections are free of adipose tissue and paraffin. If additional adipose tissue or wax is evident, the slide should be re-submerged in xylene. Use caution to minimize added exposure to xylene as this can cause shrinkage of the tissue.
   3. Rehydrate tissues in a slide jar containing 100% ethanol for 10 min. Repeat once.
   4. Move slides to a slide jar containing 95% ethanol for 10 min. Repeat once.
   5. Move slides to a slide jar containing 75% ethanol for 5 min.
   6. Move slides to a slide jar containing 50% ethanol for 5 min.
   7. Boil 10 mM sodium citrate solution (pH 6.0) in a hot plate or a microwave and pour into Coplin jar.
      CAUTION: Carefully monitor solution while heating and take care to avoid boil over. Container will be very hot. Use protection to avoid burns.
   8. Slowly place slides into Coplin jar, ensuring complete submersion, and incubate at 100 °C for 10 min to retrieve epitopes. Remove and carefully dry slides.
   9. Draw a barrier around tissue with a hydrophobic marker.Add enough endogenous enzyme blocker (hydrogen peroxide and sodium azide, available commercially) to cover tissue and incubate for 10 min.
   10. Submerge slides in 1x phosphate buffered saline (PBS) for 10 min.
   11. Add enough 10% donkey serum in 1x PBS to cover tissue and incubate for 1 h at room temperature in a humidified chamber.
       NOTE: The experiment can be paused here by storing the slides in the humidified chamber at 4 °C overnight. While donkey serum was found to yield optimal staining results, the investigator may wish to test different sera such as bovine serum albumin (BSA), goat serum or horse serum to determine optimal results. If using BSA, the investigator should prepare this fresh before use.
   12. Decant excess blocker from slides and add properly diluted primary antibody in 1% serum directly to slides ensuring that the tissue is evenly covered in solution. Incubate for 30 min at room temperature in a humidified chamber.
       NOTE: This step may be continued for longer time durations with humid chamber stored at 4 °C. Make sure to include proper negative control slides (e.g. a slide with no primary antibody, known antigen-negative tissue, etc.) at this step.
   13. Carefully wash slides by flowing about 1 mL of diH₂O over tissue and incubate in 1 change of 1x PBS for 5 min.
   14. Decant excess PBS and add enough horseradish peroxidase label to cover tissue and incubate for 30 min in a humidified chamber.
       NOTE: At this step, the investigator may wish to proceed to fluorescent staining using fluorescently conjugated secondary antibodies. If this is desired, the next steps described should be modified accordingly.
   15. Carefully wash slides by flowing about 1 mL of diH₂O over tissue and incubate in 1 change of 1x PBS for 5 min.
   16. Make diaminobenzidine (DAB) plus chromogen solution according to the manufacturer suggested ratio and mix by vortex.
       NOTE: Many ready-made kits are commercially available aside from the one used in this protocol.
   17. Decant excess buffer from slides and add 2-3 drops (10-50 µL) of chromogen stain directly to tissues and incubate for 5 min in humid chamber. Carefully wash slides by flowing about 1 mL diH₂O over tissue.
       CAUTION: DAB-chromogen is highly toxic. Avoid direct contact of stain with skin and collect flow off in hazardous waste.
2. Hematoxylin and Eosin (H&E)
   1. Following final rinse after chromogen incubation, submerge slides in Harris hematoxylin for 2-2.5 min. Rinse slides gently under tap water for 1-2 min.
      NOTE: Take care to avoid direct jet of water onto tissues.
   2. Submerge slides for 2-3 s in differentiation solution (0.25 mL of hydrochloric acid in 100 mL of 70% ethanol). Rinse slides gently under tap water for about 1-2 min.
   3. Submerge slides in blue agent (4.5 mg calcium carbonate in 100 mL tap water, pH adjusted to 9.4) for 60 s. Rinse slides in 95% ethanol for 30 s.
   4. Submerge slides in alcoholic eosin Y for 2-3 min.
   5. Dehydrate tissue in 2 changes of 95% ethanol for 1 min each.
   6. Dehydrate tissue in 1 change of 100% ethanol for 1 min.
   7. Carefully dry slides with a lint-free wipe. Apply 1 drop (about 100-200 µL) of synthetic, non-aqueous, resin-based mounting media to slide and apply coverslip.
      NOTE: If a different staining method was elected, a different mounting media may be required. For instance, if the investigator chose a fluorescent-conjugated secondary antibody, an aqueous mount would be more ideal.
   8. Allow slides to set overnight at room temperature.
3. Picrosirius Red (PSR) Staining
   1. Select slides to be stained and follow immunohistochemistry protocol through step 6 (50% ethanol soak).
   2. Submerge slides in PSR stain for 1 h.
   3. Submerge slides in 0.5% acetic acid for 1-2 s, twice to differentiate stain.
   4. Dehydrate tissues in 2 changes of 95% ethanol for 1 min each.
   5. Dehydrate tissues in 1 change of 100% ethanol for 1 min.
   6. Clear slides by briefly submerging in 100% xylene for about 3-4 s.
   7. Carefully dry slides with a lint-free wipe. Apply 1 drop (about 100-200 µL) of synthetic, non-aqueous, resin-based mounting media to slide and apply coverslip.

3. Sample Analysis

1. Ductal Analysis
   1. Select slides stained for α-SMA. Using a light microscope fitted with a camera-mounted objective, collect representative images at 20X magnification.
   2. Using ImageJ (NIH), distinguish and count ducts in each image. These can be enumerated manually or one may assign a numerical score to individual ducts. To assign a numerical score, open ImageJ and select Plugins. Next select Analyze and choose Cell Counter from the drop down menu. Click Initialize then highlight Type 1 to begin labeling ducts. When finished, select Results to view the ductal count.
      NOTE: Take care to verify structures are ductal and not vascular.
   3. In 'Set Measurement' options, check 'Perimeter' and 'Area".
   4. Using the freehand polygon tool, draw a line around each duct at the myoepithelial compartment (visibility aided by α-SMA stain). Select 'Measure' and record the perimeter and area.
   5. Again using the freehand polygon tool, draw a line along the apical side of the ductal epithelium within the interior of the lumen. Select 'Measure' and record the perimeter and area.
   6. Subtract the perimeter of the luminal compartment from the perimeter of the myoepithelial compartment in order to obtain the circumference of the ductal epithelium. Subtract the area of the luminal compartment from the area of the myoepithelial compartment to obtain the area of the ductal epithelium.
2. Immunohistochemistry Analysis
   1. Upload brightfield images to an analytical software, such as ImageJ or FIJI Suite.
      NOTE: This protocol outlines the steps for staining analysis using ImageJ and the IHC Toolbox plugin.
   2. Open the IHC Toolbox from the Plugin menu. In the "Select Model" combo box that opens, select appropriate stain (i.e. H-DAB, PSR, etc.). Select the "Color" option to isolate the stain. This will open a result window.
      NOTE: This method is appropriate if a protein of interest is located in the extracellular or cytosolic spaces, or is bound to the plasma membrane. If the protein of interest is nuclear, selecting "Nuclei" will prompt the plugin to analyze the image for positively stained nuclei.
   3. Use the Color Chooser Slide to ensure proper isolation of the stain without background inclusion or excessive stain exclusion.
      NOTE: If the automatic mode is unable to detect the stain or does not result in acceptable images, draw a square Region of Interest (ROI) over an identified stained region. On the IHC Toolbox window, select "Train" to direct the plugin to the appropriate target.
   4. Convert the image to 16-bit. Go to Image → Type → 16 bit.
   5. Threshold the image. Go to Image → Adjust → Auto Threshold.
   6. Set the image measurement scale by drawing a line ROI directly over the scale bar in the image then assigning the length. To set the scale bar, go to Analyze → Set Scale. Input the number of pixels per the desired unit of length (e.g. 120 pixels per 50 µm).
   7. Measure the resultant area and mean grey value. Go to Analyze → Set Measurements and collect the measurement by clicking Analyze → Measure.

Results

Female mice have 5 pairs of mammary glands. Specifically, there is one pair of cervical glands (#1), two pairs of thoracic glands (#2 and #3), one pair of abdominal glands (#4), and 1 pair of inguinal glands (#5) (Figure 1A). Here, we isolated the #4 glands as they are readily identifiable. In some circumstances, both #4 and #5 glands were isolated together as distinction between the two was difficult. To isolate intact #4 abdominal mammary g...

Discussion

In the paper, we have described a technique to isolate intact mouse mammary glands for downstream histological analyses of ECM expression and ductal morphology. With respect to analyses of ductal morphology, this methodology enables the rapid investigation of ductal architecture based off of stained histological sections. Other methods of ductal analyses rely on injections of dyes to enable visualization of the ductal tree, methods which may be technically challenging and time consuming.

In br...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Light Microscope	Olympus	BX43
Microscope Camera	Olympus	DP73
Image Analysis Software	Olympus	cellSens Entry software
	NIH	ImageJ
3.5x-R Surgical Micro Loupes	Rose Micro Solutions		Magnification at researcher's preference
Mayo Scissors	Medline	DYND04035
Staining Rack	Fisher Scientific	121
Staining Dish	Fisher Scientific	112
Coplin Jars	Fisher Scientific	19-4
Glass coverslips	Fisher Scientific	12-550-15	Size appropriate for tissue
IHC EnVision+ Kit (HRP, Mouse, DAB+)	Dako	K400611-2
Picrosirius Red Kit	Abcam	AB150681
Eosin Y, alcoholic	Sigma-Aldrich	HT110132
Harris Hematoxylin	Sigma-Aldrich	HHS16
Donkey Serum	EMD Millipore	S30
10% Neutral Buffered Formalin	Sigma-Aldrich	HT501128
Xylenes, Reagent Grade	Sigma-Aldrich	214736
Ethanol, 200 proof	Sigma-Aldrich	792780	suitable for molecular biology
Phosphate Buffered Saline, 1x	Gibco	10010023
Sodium Citrate	Fisher Scientific	S279-500
Calcium Carbonate	Sigma-Aldrich	202932
Permanent Mounting Medium	Dako	S1964
Eukitt's Mounting Medium	Sigma-Aldrich	3989
Fibronectin antibody	Abcam	AB23750
Tenascin-C antibody	Abcam	AB108930
Alpha Smooth Muscle Actin antibody	Abcam	AB124964
Dako Envision Dual Link System HRP	Dako	K4065