Rat Mammary Epithelial Cell Transplantation into the Interscapular White Fat Pad

Summary

This article describes a transplantation method to graft donor rat mammary epithelial cells into the interscapular white fat pad of recipient animals. This method can be used to examine host and/or donor effects on mammary epithelium development and eliminates the need for pre-clearing, thereby extending the usefulness of this technique.

Abstract

As early as the 1970s, researchers have successfully transplanted mammary epithelial cells into the interscapular white fat pad of rats. Grafting mammary epithelium using transplantation techniques takes advantage of the hormonal environment provided by the adolescent rodent host. These studies are ideally suited to explore the impact of various biological manipulations on mammary gland development and dissect many aspects of mammary gland biology. A common, but limiting, feature is that transplanted epithelial cells are strongly influenced by the surrounding stroma and outcompeted by endogenous epithelium; to utilize native mammary tissue, the abdominal-inguinal white fat pad must be cleared to remove host mammary epithelium prior to the transplantation. A major obstacle when using the rat model organism is that clearing the developing mammary tree in post-weaned rats is not efficient. When transplanted into gland-free fat pads, donor epithelial cells can repopulate the cleared host fat pad and form a functional mammary gland. The interscapular fat pad is an alternative location for these grafts. A major advantage is that it lacks ductal structures yet provides the normal stroma that is necessary to promote epithelial outgrowth and is easily accessible in the rat. Another major advantage of this technique is that it is minimally invasive, because it eliminates the need to cauterize and remove the growing endogenous mammary tree. Additionally, the interscapular fat pad contains a medial blood vessel that can be used to separate sites for grafting. Because the endogenous glands remain intact, this technique can also be used for studies comparing the endogenous mammary gland to the transplanted gland. This paper describes the method of mammary epithelial cell transplantation into the interscapular white fat pad of rats.

Introduction

Postnatal mammary gland development and ductal morphogenesis are processes largely influenced by hormonal signaling at the onset of puberty. In mice and rats, commonly used model organisms of mammary gland biology, this process begins around 3 weeks of age, where rapid proliferation and differentiation result in the formation of the mature parenchyma. The mature mammary gland can undergo numerous rounds of expansion and involution, a property that has been under investigation since the early 20^th century. Within the context of hyperproliferation and cancer development, mammary gland transplantation techniques were developed in the 1950s¹, and enhanced by the quantitative methodology contributed by Gould et al. in 1977^2,3,4. Refinement of the transplantation technique in rodents has contributed to major advances in understanding normal mammary gland biology that are still widely used to study the effect of various treatments and genetic manipulation on normal mammary gland development and disease states.

Many hypotheses have been generated and subsequently tested using mammary gland transplantation, first described by DeOme et al. in 1959¹. Experiments across several decades showed the propensity of ductal tissue excised from donor mammary glands to repopulate the entire fat pad^5,6,7 and indicated that a critical component of mammary gland development resides in these epithelial structures. Later studies in mice showed that a single mammary stem cell can repopulate a cleared fat pad and contributed to the discovery of a single, common progenitor of basal and luminal mammary epithelial cells^8,9,10. In line with these conclusions, it has been suggested that transplantation increases the pool of cells with multilineage-repopulating potential as a result of plasticity, allowing the grafted cells to grow a functional mammary gland^{7,10,11,12,13}. Importantly, the use of transplantation techniques in rodents overcomes the limitations of cell culture-induced abnormalities¹⁴ and often provides results in just a matter of weeks.

While the procedure was originally described in the context of preneoplastic lesions in mice, it was soon expanded to rats and used in conjunction with the carcinogen treatment to establish multiplicity as a measure of cancer susceptibility¹⁵, but the popularity of transplantation techniques has followed the development of genetic tools for each species. Although mouse studies incorporating transplantation have contributed many translational findings, the parenchyma of the rat mammary gland resembles the human more closely^16,17 and offers distinct advantages for studying estrogen receptor-positive (ER+) breast cancer. Mammary tumors are inducible in both species, but they differ in terms of hormone sensitivity and gene expression profiles. A primary difference is that rat mammary tumors express and depend on the function of ovarian and pituitary hormone receptors, namely, estrogen and progesterone (PR), similar to the luminal-A subtype of human breast cancer. Indeed, mammary epithelial cell transplantation, as described in this protocol, has been used to study genetic variants involved in breast cancer and determine the cellular autonomy of effects on mammary epithelial cells¹⁸.

In addition to the tumor biology, the ductal epithelium of the normal rat mammary gland exhibits a higher level of branching and is flanked by a thicker layer of stroma than the mouse. The importance of the stroma is well-documented in mammary epithelial transplantation studies. Mammary epithelium must interact with fatty stroma, and ideally its own mesenchyme, to undergo its characteristic morphogenesis^19,20. Grafting tissue into a recipient mammary gland provides an optimal environment; however, the presence of endogenous epithelium can interfere with results. Preclearing the mammary gland of endogenous epithelium is commonly performed in mouse transplantation assays and requires surgical excision of endogenous mammary tissue and/or removal of the nipple^1,21,22. Although possible, preclearing the mammary epithelium in post-weanling rats is not as widely-performed, mainly due to the ineffectiveness of clearing the growing mammary tree in post-weanling rats. Since it has been shown that regions of adipose tissue elsewhere in the body could support the growth of transplanted mammary epithelium^21,23,24, the process of preclearing can be easily avoided in rats by grafting tissue into the interscapular white fat pad.

The transplantation method described in this paper involves the injection of enzymatically dissociated mammary gland organoids (fragments of mammary ductal epithelium and other cells types capable of morphogenesis) or monodispersed cells into the interscapular fat pad in inbred, isogenic or congenic strains of laboratory rats². Because the interscapular fat pad is normally devoid of mammary tissue, it provides a suitable environment for multiple transplantation sites without the need to pre-clear endogenous epithelium. As a result, the host animal's endogenous, abdominal-inguinal mammary glands are not subject to surgical manipulation, develop normally, and cannot interfere with interpretation of results. Additionally, the intact mammary glands can be used for comparison to evaluate host versus donor effects on the mammary epithelium development and tumorigenesis^18,25. Although repopulation of the mammary gland from a single stem cell is available for mice, it has not yet been developed for rat, mainly due to the lack of availability of antibodies to select for rat mammary stem cells^25,26,27. Despite this, transplantation of monodispersed mammary epithelial cells to quantify repopulating potential can be successfully performed, and those cells will develop normally when grafted into the appropriate framework^2,3,4. While organoids are good for many purposes, monodispersed cells are required for quantitative applications, for example, to determine the number of mammary epithelial cells required for the cancer initiation following ionizing radiation treatment²⁸ or for comparing characteristics of flow cytometrically selected mammary epithelial cell populations²⁹.

To date, the procedure described here is the most robust method of performing mammary gland transplantation in the rat with an overall goal of studying mammary gland development and mechanisms underlying breast cancer development. Often, the donor and/or recipient animals are exposed to different variables before, during, or after the epithelial transplantation. Examples include single gene studies involving chemical carcinogenesis³⁰, radiation^28,31,32, genetic manipulation of host/donor genome¹⁸, and hormonal manipulation¹². A major advantage of the enzymatic dissociation described in this protocol is the opportunity to isolate epithelial organoids or monodispersed cells for complementary experiments involving flow cytometry, 3-D culture, gene editing, and more. Future applications of this technique will include additional manipulation of donor and/or host tissue with genetic engineering. For example, donor cells can be genetically altered ex vivo at any chosen genomic locus using the CRISPR-Cas9 gene editing system. Similarly, recipient rats can also be genetically altered to study the interaction between donor and recipient engineered genetic factors.

Protocol

All animals were housed and maintained in an AAALAC-approved facility, and experiments described in this protocol were approved by the MUSC Institutional Animal Care & Use Committee (IACUC). Animals for use in reciprocal transplantation should be an inbred or isogenic strain, with congenic status preferred or backcrossed for at least 6 generations.

1. Harvesting donor rat mammary gland epithelium

1. Determine the number of donor rats needed for transplantation.
   NOTE: Generally, 1 donor rat (4 weeks of age) can provide enough cells for transplantation into 4 recipient animals. Certain applications of this protocol will require additional numbers of cells, and there can be strain-specific differences in total yield.
2. Label all supplies and ensure accessible placement for the surgeon (Table 1).
3. Record the body weight of each female donor rat. Follow institutional guidelines to fully anesthetize or euthanize the donor rat. Check for the depth of anesthesia by the lack of response to the toe pinch.
4. Move the animal to the sterile surgical field. Place the animal on its back and spray the entire ventral surface with 70% ethanol.
5. Make a sagittal, X-shaped incision, allowing access to thoracic, abdominal and inguinal mammary glands. Dissect all the mammary gland tissue from the donor rat using scissors (Figure 1). Remove all visible lymph nodes.
6. Extract the mammary tissue using forceps and place in a labeled 60 mm dish on ice. Add 500 µL of serum-free DMEM/F12 media to the 60 mm dish. Adjust the placement of the mammary gland tissue so it stays completely wet.
7. Finely mince the mammary tissue using scissors. To do so, cut the tissue into pieces 1-2 mm³ in size (Figure 1C). Keep the gland on ice until all the donor tissue has been harvested (do not exceed 60 min).
   NOTE: Additional personnel can mince mammary glands and additionally prepare collagenase solution while the surgeon proceeds with tissue extractions.

2. Extract brain tissue from euthanized donors

1. Carefully turn the body of the donor animal over to place it in a prone position. Secure with pins. Spray the head and upper back with 70% ethanol.
2. Locate the base of the skull and make an incision beneath the occipital condyles. Insert sharp scissors beneath the skin and cut the skin away from the skull, including the sides of the head.
3. Use bone cutters or strong scissors to cut the skull along the midline, from the occipital to frontal bones. Keep the blade as superficial as possible and angle upward to prevent the destruction of the underlying brain tissue.
4. Peel the bone away using rongeurs or strong forceps. Insert the tool lateral to the cerebellum to break the bone on either side, exposing the bony auditory canal. Sever the connections to the meninges.
5. Gently lift the brain with curved, fine-tip forceps. Place the brain on a piece of foil and record the weight, and then immediately transfer to a 15 mL tube with an equal amount (w:v) of media, stored on ice.
   1. Optionally, use fine-tip forceps to remove the pituitary gland (located beneath the brain) for additional use in the transplant procedure, if needed.
6. Use a mechanical homogenizer to disrupt the tissue. Homogenize the brain for 10-15 s on low speed. Let the mixture sit on ice for at least 1 min, and then homogenize again. Homogenization is sufficient when the final mixture is free of large pieces.
7. Filter the homogenate by passing it through a 100 µm filter. Keep the filtrate on ice until use (less than 4 h).

3. Digestion and processing of mammary gland extracts

1. Thaw or warm reagents as indicated (Table 1). Follow the appropriate steps for recovering organoids or monodispersed cells.
2. Prepare 10 mL of serum-free digestion media (without collagenase) for every donor animal (Supplemental File 1).
   NOTE: The volume of digestion media used can be adjusted (scaled up or down) to accommodate the grouped tissue, if applicable.
   1. For organoids skip to 3.3.
   2. For monodispersed cells, prepare fresh Monodispersion Mixture and Inactivation Solution (Supplemental File 2, Supplemental File 3) in addition to the serum-free collagenase digestion media. Proceed to step 3.3.
3. When all the mammary tissue from donors has been extracted and minced, add the collagenase enzyme to the warm (or room-temperature) digestion media (Supplemental File 1). Mix by inverting.
4. Pass the collagenase digestion media through a 20 µm filter. Dispense 10 mL of filtered media in the labeled 50 mL tubes for digestion.
5. Use a new 1,000 µL pipette tip for each sample and transfer the minced donor tissue from each 60 mm dish to the collagenase digestion tube. Cut 1 cm off the end of a 1,000 µL tip and manually place it on the device before use. Gently mix the minced tissue by pipetting up and down 1-2 times.
   NOTE: If the tissue is difficult to pipette during transfer, use a small amount of the collagenase digestion media from the 50 mL tube, and then transfer it back.
6. Place the samples in the horizontal position in a shaking incubator and allow the samples to digest 1.5-2 h at 37 °C, 200-220 rpm.
   1. For organoids skip to 3.7.
   2. For monodispersed cells add DNase I (0.2 µg/mL) to the mixture for the last 10 min of the digestion. Incubate as before, with vigorous shaking. Proceed to step 3.7.
7. When the tissue is fully digested, pellet the suspension using cold centrifugation (4 °C) for 10 min at 1,200 x g (Figure 1D).
   NOTE: Keep tubes on ice between every step to increase viability of cells.
8. Ensure that a pellet has formed, and then carefully pour off the supernatant and fat layer. Gently resuspend the pellet in 10 mL of fresh DMEM/F12 media.
9. Briefly spin at 68 x g for approximately 10 s. The length of the spin (but not the speed) may be increased if there is no clear separation of cells. Visually inspect the pellet before proceeding (Figure 1D).
10. Carefully remove the supernatant, leaving behind a small volume. Proceed to the next step, based on whether organoids or monodispersed cells are needed.
    NOTE It is important to leave a small volume of media in the tube because the pellet will be very loose. The residual volume of media will be diluted through wash steps.
    1. For organoids, add another 10 mL volume of DMEM/F12 media and repeat the wash/spin. After the second wash, resuspend the cells in a smaller volume (1-2 mL) of DMEM/F12 media for filtration. Proceed to step 3.11.
    2. For monodispersed cells, dissolve in 2 mL of pre-warmed HBSS with 0.025% (w/v) Trypsin and 6.8 mM EDTA. Digest for 3-5 min at 37 °C. Inactivate immediately.
       1. Add 4 mL of DMEM/F12 with 10% FBS to stop inactivate the trypsin.
       2. Spin the cells at 270 x g for 5 min, and then resuspend in DMEM/F12 with 10% FBS again. Proceed to step 3.11 to filter the cells.
11. Filter the cells using a 40 µm cell strainer placed in a new 50 mL tube. Pre-wet the strainer by pipetting 1 mL of the same base medium used to suspend the cells, and then pass the cell suspension through the filter using a pipette to collect ductal fragments/organoids.
    NOTE: The approximate yield of filtered epithelium from the mammary gland tissue of a single, 4-week-old donor rat is 1 x 10⁶ cells.
    1. For organoids, discard the filtrate. Mammary organoids will remain inside the basket of the cell strainer and smaller, unwanted cells will be eliminated. Invert the cell strainer over a new 50 mL tube, and rinse with any volume necessary to collect the cells. Proceed to step 3.12.
    2. For monodispersed cells, discard the cell strainer and keep the filtrate because the mammary epithelial cells will pass through the filter, along with smaller stromal and immune cells. Rinse the tube that was used for the digestion/centrifugation with another volume of DMEM/F12 with 10% FBS and pass through the same cell strainer. Pellet the monodispersed cells by centrifugation at 1,200 x g for 5 min. Proceed to step 3.12.
       NOTE: The resuspended cells are ready for subsequent applications.
12. Pulse spin the solution and ensure the formation of a cell pellet before proceeding to the next step. Pulse spin again, if necessary.
13. Carefully remove the supernatant. Resuspend the pellet in a small volume (1,000-2,000 µL of DMEM/F12) to concentrate the cells for counting.
14. Count cells and dilute if needed. Resuspend the desired number of cells for transplantation in 20 µL of DMEM/F12 media for each animal. Always keep cells on ice.
    NOTE: Donor cell counts in the range of 1 x 10⁵ - 1 x 10⁶ cells will be required for each graft site based on the endpoint of the study. For example, carcinogenesis experiments often require a greater number of transplanted cells, relative to other applications. The number of cells needed must be experimentally determined for each strain. The procedure described in this protocol utilized 250,000 donor cells in 20 µL media per graft site, prior to mixing with brain homogenate, as described in step 3.16.
15. Prepare any aliquot(s) of cells needed for other experiments (e.g., FACS isolation^{3,26,27,30,33}).
    1. For organoids, proceed to step 3.16.
    2. For monodispersed cells, quantify the viable cells using Trypan or methylene blue staining, and then proceed.
16. Prepare single batches of donor material for all the transplant recipients. Combine equal volumes of the cell suspension (20 µL) with 50% brain homogenate (20 µL) for every site of transplantation.
    NOTE: A total of 40 µL per site will be injected at each site, but it is recommended to include a minimum of 25% extra volume for waste.
17. Immediately proceed to transplantation or freeze cells for the transplantation later (potential stopping point).
    NOTE: Frozen cells have not been tested with this protocol and will require optimization in advance of generating a cohort of animals for transplantation experiments. It is strongly recommended to transplant fresh cells.

4. Transplantation procedure (recipient rats 4-5 weeks of age)

1. Weigh each recipient rat and calculate the correct dose of approved analgesic that will be used in the procedure.
   NOTE: Body weights can be measured up to 24 h in advance of the procedure. Follow institutional guidelines to restrain or briefly anesthetize each animal for the duration of shaving.
2. Shave the surgical area on each animal using electric clippers. Identify the base of the skull and start of the vertebral column. Approximately one-third of the way down the spine, shave a 3 cm x 2 cm area on the upper thoracic portion of the back.
   NOTE: Shaving can also be performed under anesthesia on the day of transplantation but must be performed outside the sterile field. Return the rat to its home cage until it is needed.
3. Locate all of the supplies needed for transplantation (Table 1).
4. Evaluate the Laboratory Animal Anesthesia System before use. Top off any fluids and replace any tanks or parts that are needed. Ensure the gas line to the anesthesia chamber is open and all peripheral lines are closed so isoflurane anesthesia and oxygen may freely flow to the animal once it is placed in the chamber.
5. Warm heating pads to support the body temperature of recipient animals.
6. Generate the sterile field that will be used for surgery. Arrange the supplies as described in Table 1.
7. Administer preoperative analgesic to recipient rats as indicated by institutionally-approved animal care protocol.
8. Flush the Hamilton syringes with sterile DMEM/F12 media to prevent loss of cells. Ensure the needle is secured to the body of the syringe, insert the needle into the liquid, and draw back the plunger. Fill to the maximum volume. Press the plunger down and expel the contents into a waste collection tube. Repeat 3-5 times.
9. Load the entire volume of donor material (prepared in step 3.16) into a separate syringe for each condition (e.g., control, treated, wildtype, knockout, etc.). Insert the tip of the needle into the liquid, draw back the plunger and keep the needle beneath the surface of the mixture as the volume in the tube decreases.
   NOTE: Include at least 10% extra volume in each syringe. Do not dispose of the remaining mixture close the tube and keep it on ice in case more is needed.
10. Invert the syringe after it is fully loaded and press the plunger slightly to remove air bubbles at the tip of the needle. Proceed to the next step when everything is prepared.
    NOTE: Make sure the tip of the needle never touches any other surface, even within the sterile field. It is helpful to rest the body of the syringe over a small container of ice to promote viability of the cells.
11. Place the recipient animal in the anesthesia chamber and turn on the machine.
12. When the animal is fully relaxed (does not react to tapping or gentle movement of the chamber), direct the anesthesia to the nose cone and transfer the animal to the sterile field.
    NOTE: Extended duration of anesthesia is not well-tolerated by rats. Complete the procedure for each animal in 10 min or less.
13. Place the animal in a prone position (on its stomach) so the back of the head and the upper spine is accessible.
    NOTE: Ensure adequate heat support for the animal all times and regularly assess the depth of anesthesia using a firm toe pinch.
14. Optionally, apply ophthalmic vet ointment to prevent drying of the eyes.
15. Clean the freshly-shaved area to remove excess hair. Use a circular motion and apply 70% ethanol (or another reagent, per institutional guidelines) to the skin, followed by an antiseptic (such as iodine), and repeat. Place a towel drape over the animal so only the region for the shaved area is exposed.
16. Ensure the animal remains unresponsive to deep stimuli with a firm toe pinch, and then proceed to the next step.
17. Make a small (2 cm) interscapular incision using a sharp surgical blade.
    NOTE: The cut must be superficial, as the fat pad is located just beneath the skin.
18. Locate the medial blood vessel for orientation (Figure 2B).
19. Lift the skin on one side of the incision using forceps and hold it away from the fat pad while the transplant is performed (Figure 2B). Insert the needle into the graft site.
    1. Optionally, move the tip of the needle inside the tissue and create a small pocket to collect the cells. Use a small, repetitive motion. Do not remove the needle.
       NOTE: This step is recommended for first-time users of the protocol. Use extreme caution when creating a pocket, as the interscapular fat pad tissue is very delicate.
20. Carefully inject 40 µL of the cell mixture into the interscapular fat pad tissue. Remove the needle slowly.
21. Hold the tissue in place and allow the transplanted cells to settle for 3-5 s. Use an additional pair of forceps, if needed.
22. Remove the needle. Repeat the injection procedure (steps 4.18-4.21) at the second site of transplantation.
    NOTE: The epithelium from one donor group can be injected into the same side of the fat pad in every animal, or alternating sides to prevent batch effects from the hand-dominance of the surgeon.
23. Close the surgical wound using wound clips or sutures, and then discontinue anesthesia.
24. Provide post-operative analgesic as indicated by institutionally-approved protocol.
25. Immediately move the animal to a recovery cage with the heat support. Monitor for signs of distress such as bleeding from the incision or trouble breathing.
    NOTE: The animal should fully recover within 5-10 min. Refer to institutional guidelines for returning animals to the colony and post-operative monitoring after survival procedures.
26. Optionally, perform carcinogenesis studies at the graft site(s) by administering carcinogens to the recipient rats 3-4 weeks after transplantation.
    NOTE: Typically, rat mammary carcinogenesis is performed using a chemical carcinogen treatment at 50-57 days of age. This treatment dictates the age of the transplant surgery (which must be done between 29-36 days of age) to allow enough time for the grafted cells to initiate growth of the mammary gland.

5. Assessment of epithelial outgrowth

1. Monitor the estrus cycle of rats through daily vaginal lavage and examine the cytology on a microscope slide. Begin 8-12 days before the endpoint of the study. Sacrifice all rats in the same stage. This is an optional step.
   NOTE: The rat estrus cycle is 4-5 days. Allowing the animal to go through 1-2 full cycles will facilitate interpretation, as lavage slides from previous cycles can be used for comparison.
2. Sacrifice transplant recipient rats 6-8 weeks after transplantation, per institutional guidelines.
   NOTE: Outgrowth is usually detectable 3-6 weeks after transplantation, but additional time may be required.
3. Place the animal in a prone position and clean the body with 70% ethanol. Lift the skin with forceps and make an incision along the vertebral column to expose the interscapular fat pad. Dissect the skin away from the tissue so the majority of the fat pad is visible.
4. Identify the medial blood vessel that separates the graft sites in the interscapular fat pad. Excise the entire pad as a single piece of tissue or cut along the blood vessel and remove sides individually.
5. Place the tissue on a positively-charged microscope slide for whole mount. Use 2 pairs of blunt forceps and gently spread the tissue to restore its original conformation on the slide.
   NOTE: Rat mammary tissue is extremely delicate. The edges of the tissue may curl under itself. Always handle with care and hold in place until the tissue adheres to the slide (a few seconds).
6. Whole-mount at least one of the endogenous abdominal-inguinal mammary glands (with lymph nodes for orientation) for comparison.
7. Place the slides in 70% ethanol for 7-10 days to defat the tissue. Replenish ethanol as often as necessary to ensure the tissue does not dry out.
8. Prepare alum-carmine stain and process the slides when the tissue is sufficiently opaque. Allow the stain to cool before usage (Supplemental File 4).
   NOTE: The stain can be prepared up to one day in advance of the fixation and rehydration steps. The solution can be stored at 4 °C and has limited potential for reuse.
9. Fix the tissue by placing the slides in 25% glacial acetic acid : 75% ethanol for 60 min.
   1. Rehydrate the tissue through a series of 3 washes in a series of decreasing concentration of ethanol: 70% ethanol for 15 min, 50% ethanol for 5 min and dH₂O for 5 min.
10. Stain with alum carmine for 4-8 days. Check the back of the slides each day to determine if the stain has fully penetrated the tissue. Proceed to the next step when the staining is complete.
    NOTE: Staining is complete when the thickest parts of the gland have a purple hue and no longer appear white.
11. Destain and dehydrate the tissue by transferring the slides through a series of increasing concentration of ethanol: 70% ethanol for 30 min, 95% ethanol for 30 min and 100% ethanol for 30 min.
12. Place dehydrated slides in xylene for 3+ days to clear the tissue. Transfer to mineral oil for long term storage.
13. After the slides have cleared, use low-powered light microscopy or high-resolution digital photography to acquire images of the slides for analysis. Ensure image acquisition parameters are consistent for all slides.
    NOTE: Epithelial outgrowth must be clearly distinguishable.
14. Treat the presence of outgrowth as a binary outcome.
15. Calculate the mean number of transplanted epithelial cells that produced ≥1 mammary outgrowth in 50% of graft sites using the acquired images. Quantification other physical features, as needed.

Results

Donor and recipient mammary glands
The steps to isolate and prepare rat mammary epithelial cells for transplantation are shown in Figure 1A. At 4 weeks of age, the endogenous mammary gland of the donor rat has begun maturation and epithelium can be visualized on whole mounted slides stained with alum carmine (Figure 1B). One donor rat at this age ...

Discussion

This protocol describes a mammary epithelial cell transplantation technique optimized for working with rats. Isolated mammary epithelial organoids from donor rats (3-5 weeks of age) are grafted into the interscapular white fat pad of recipient rats (also 3-5 weeks of age). Results can be interpreted as little as 4-6 weeks later, using light microscopy to examine the grafted tissue; however, the optimal amount of time between transplantation and sacrifice must be determined prior to implementing a full experiment. If too ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.2 µM syringe filters	Fisher Scientific	09-715G	sterile-filtering collagenase digestion media
1.5 - 2.0 mL microcentrifuge tubes (sterile)	Fisher Scientific	05-408-129	containing resuspended cells and/or brain homogenate mixture
100 µM cell strainers	Corning	431752	filtering brain homogenate
100 uL gastight syringes with 25 gauge needles	Hamilton	81001 & 90525	For injecting graft mixture into recipient animals (1 per donor genotype/condition)
1000 uL pipette tips + pipette	-	-	transferring cells/mixtures/tissue
15 mL polypropylene tube	Falcon (Corning)	352196	brain homogenate mixture storage, or cell : homogenate mixture for transplantation
40 µM cell strainers	Corning	431750	filtering organoids after washing the cell pellet
50 mL polypropylene tubes	Fisher Scientific	05-539-6	for collagenase digestion of donor mammary gland tissue
60 mm dishes	Thermo Scientific	130181	for mincing tissue
Alum Potassium Sulfate	Sigma-Aldrich	243361/237086	staining mammary gland whole mount slides
Anesthesia vaporizer for veterinary use	-	-	follow institutional protocol
Beta-dine or iodine	-	-
Borosilicate glass culture tube for homogenization	Fisher Scientific	14-961-26	for homogenization of brain (use appropriate tube for homogenizer)
Carmine	Sigma-Aldrich	C6152/1022	staining mammary gland whole mount slides
Cell counting apparatus	-	-
Clean animal cages for recovery	-	-	follow institutional protocol
Collagenase Type 3	Worthington Biochemical Corp.	LS004183	enzymatic digestion of minced mammary gland tissue from donor rats
deionized water	-	-	for chemical solutions
DMEM/F12	GIBCO	11320033	for mincing tissue, collagenase digestion media and resuspending epithelial cell mixtures
EDTA	-	-	monodispersion mixture
Ethanol, 200 Proof	Decon Labs	2705/2701	mammary whole mount slide fixative, mammary whole mount slide washes, cleaning surgical incision sites (diluted)
Fetal Bovine Serum (FBS)	Hyclone	-	inactivation solution
Gauze	-	-
Glacial acetic acid	Fisher Scientific	A38-212	use for mammary whole mount slide fixative (1:4 glacial acetic acid in 100% ethanol)
HBSS	GIBCO	-	monodispersion mixture
Heating pads	-	-	follow institutional protocol
Ice buckets (x2)	-	-
Incubator with orbital rotation	-	-	must be capable of maintaining 37°C, shaking at 220-225 RPM (for collagenase digestion of mammary tissue)
Isoflurane anesthesia	-	-	follow institutional protocol
Light microscope or digital camera	-	-	visualizing whole mounted mammary epithelium and/or acquiring images
Mechanical homogenizer	Fisher Scientific	-	TissueMiser or alternative models
Mineral oil, pure	Sigma-Aldrich/ ACROS Organics	8042-47-5	long-term storage of cleared mammary gland whole mounts
Oxygen tanks for anesthesia vaporizer	-	-	follow institutional protocol
Paper towels or delicate task wipes	-	-
Positively-charged microscope slides	Thermo Scientific	P4981-001	mammary gland tissue whole mounts
Postoperative analgesic	-	-	Institutional protocol
Scale			body weight measurements of animals, proper dosing of pain medication
Shaver	-	-	electric clippers, or other
Staining jars	-	-	minimum of 1 per chemical wash, size appropriate for the number of slides, glass preferred
Sterile field drapes	IMCO	4410-IMC	used during transplantation
Sterile scissors and forceps x3 (autoclaved)	-	-	autoclave surgical tools used for donors and recipients
Syringes: 5 mL (or greater)	-	-	for sterile filtration of collagenase digestion media
Trypsin	Worthington		monodispersion mixture
Waste collection receptacle for liquids (poured or aspirated)	-	-
Wound clip applier, clips, and removal tool	Fine Science Tools	12020-00	Closing the skin incision over the interscapular white pad pad
Xylenes	Fisher Scientific	X3S-4	clearing mammary gland whole mount slides after staining