Transplantation Into the Mouse Ovarian Fat Pad

All the in vivo work described is approved by the Cornell University Institutional Animal Care and Use Committee.

1. Sample Preparations for Ovarian Fat Pad Assays

1. Isolation and Culture of Primary Tubal Epithelium (TE) Cells
   1. Euthanize donor 6- to 8-week-old virgin β-actin-Enhanced Green Fluorescent Protein (EFGP) or β-actin-Discosoma Red Fluorescent Protein (DsRed) mice by CO₂ administration followed by cervical dislocation. Verify successful euthanasia by a toe pinch.
   2. Place an individual mouse on an absorbent drape, ventral side up, and then wipe the abdomen with 70% ethanol. Open the skin by making a lateral incision at the body midline and with fingertips pull the skin above and below the incision toward the head and tail of the mouse.
   3. Hold the peritoneum using blunt forceps and make a cut with fine scissors to open the body cavity. Gently push the coil of the intestine aside and locate the reproductive organs.
   4. With fine forceps pick up one uterine horn and cut 0.5 cm above the point where the uterine horns separate. While holding the uterine horn, dissect away connective tissue attached to the uterine horn, uterine tube, ovary and ovarian fat pad. Place dissected reproductive tract in a dish with 6 ml sterile Phosphate Buffered Saline (PBS). Proceed with the second reproductive tract.
   5. Transfer the dish to a Biosafety cabinet and wash the tissues 3 times in 6 ml sterile PBS. Continue the work under a dissection microscope located in the Biosafety cabinet. Grab the uterine horn with fine tweezers and disconnect the uterine horn from the uterine tube by severing at the utero-tubal junction. Cut out the ovarian bursa surrounding the ovary by using a 25 G needle.
      Note: After the ovarian bursa has been removed, the dissection is complete and the individual uterine tube remains in the PBS solution.
   6. Transfer a single uterine tube in a 50 µl drop of PBS to a 3.5 cm dish and mince into 0.1 mm pieces using 28 gauge needles. Transfer to 200 µl digestion buffer 1 (Dulbecco's Modified Eagle's Medium (DMEM) F12 (Ham's) medium supplemented with 300 units ml^-1 Collagenase and 100 units ml^-1 Hyaluronidase) and incubate for 1 hr at 37 °C in a 5% CO₂ incubator.
   7. After the incubation, add 1 ml 0.25% Trypsin-Ethylenediaminetetraacetic acid (EDTA) and suspend the solution with the help of a 1 ml pipette tip (aka blue tip) 20 times for 3 minutes. Add 5 ml + 4 °C DMEM/F12 (Ham's) medium containing 2% fetal bovine serum.
   8. Collect tissues by centrifugation (600 rcf, 5 min, room temperature (RT)), and add 1 ml digestion buffer 2 (DMEM F12 (Ham's) medium supplemented with 7 mg ml^-1 Dispase II and 10 µg ml^-1 Deoxyribonuclease I (DNase I)). Suspend tissue pellet 20 times with help of a 1 ml pipette tip.
   9. Add 5 ml serum free complete mouse tubal epithelium growth medium (M-TE-GM, Table 1).
   10. Collect cells by centrifugation (600 rcf, 5 min, RT). Add 1 ml M-TE-GM, suspend and count cells by hemocytometer. Seed 1 x 10⁵ cells in 1 ml per well in a 24-well low attachment plate and incubate in M-TE-GM at 37°C in a 5% CO₂ incubator for 4 days.
   11. Change the medium every day.
   12. Prepare single cell suspensions of cultured primary suspension TE cells from β-actin-EGFP or β-actin-DsRed mice.
       1. Collect TE suspension cultures from 24-well low attachment plates. Centrifuge (600 rcf, 5 min, RT), and suspend cell pellets with 1 ml 0.25% Trypsin-EDTA. Incubate for 10 min at 37°C in a 5% CO₂ incubator.
       2. Stop trypsin activity by adding 5 ml DMEM/F12 (Ham's) medium containing 5% fetal bovine serum. Collect cell pellets by centrifugation (600 rcf, 5 min, RT).
   13. Wash cell pellets twice with 4 ml of cold PBS (4°C), add 1 ml PBS per cell pellet, and suspend. Count cells by hemocytometer and transfer to 1.7 ml centrifugation tubes. Working on ice, add 1 x 10⁵ cells to 10 µl PBS, then add 10 µl basement membrane matrix. Suspend and transport on ice to the animal room.
       Note: Perform all dissection and tissue culture work in a Biosafety cabinet under sterile conditions.
2. Preparation of Immortalized Human Cell Line
   1. Separately grow lentivirus-EGFP and -mCherry labeled human immortalized ovarian surface epithelium cell lineHIO118 until 80 – 90% confluency on 10 cm dishes at 37°C in a 5% CO₂ incubator.
   2. Wash dishes twice with 10 ml PBS, add 1 ml 0.25% Trypsin-EDTA per dish and incubate for 10 min at 37°C in a 5% CO₂ incubator. Stop the reaction by adding 10 ml DMEM/F12 (Ham's) medium containing 5% fetal bovine serum. Collect cell pellets by centrifugation (600 rcf, 5 min, RT).
   3. Wash cell pellets twice with 10 ml of cold PBS (4°C), add 1 ml PBS per cell pellet, and suspend. Count cells by hemocytometer and transfer to 1.7 ml centrifugation tubes.
   4. Working on ice, add 1.0 x 10⁶ Lenti-mCherry labeled cells + 1.0 x 10⁶ Lenti-EGFP labeled cells to 10 µl PBS. Add 10 µl basement membrane matrix. Suspend and transport on ice to the animal room.
3. Preparation of the Uterine Tube
   1. Dissect individual uterine tubes from 6- to 8-week old virgin β-actin-DsRed mice in PBS as described in steps 1.1.1-1.1.5. Transfer single uterine tubes into a 200 µl drop of PBS under a dissection microscope. Gently uncoil uterine tubes with the help of tweezers and 28 gauge needles by removing the mesosalpinx, a portion of the broad ligament which supports the segments of the uterine tube. Place single uncoiled uterine tubes in a drop of 200 µl ice-cold PBS until the recipient's ovarian fat pad is exposed and prepared to receive the transplant.

2. Ovarian Fat Pad Transplantation

Note: Disinfect instruments between each animal surgery. Prepare and arrange all equipment in advance. Dorsal transplantations to the right ovarian fat pad are preferential as this avoids the spleen. However, recipients may have transplants in both ovarian fat pads.

1. Use syngeneic mice or severe combined immunodeficiency (SCID/NCr) mice as recipients of primary cells. For human cells, such as HIO118 cells, use Nod/SCID/gamma (NSG) mice or SCID mice.
2. Anesthetize recipient mouse with a single intra-peritoneal injection of 2,2,2-Tribromoethanol at a dosage of 0.15 ml/10 g body weight. Place the animal on a heating pad, in the animal hood and confirm proper anesthetization by loss of pedal reflex (toe pinch). Apply vet ointment on eyes to prevent dryness while the animal is under anesthesia. Inject the animal subcutaneously with the analgesic Ketoprofen at a dosage of 4 mg/ g body weight to treat for post-surgical pain.
   Note: As an alternative, use isoflurane for anesthesia. Place the animal in the induction chamber and adjust the oxygen flowmeter to 0.8-1.5 L/min and the isoflurane vaporizer to 2-3%. Remove the anesthetized animal from the chamber, let it breathe isoflurane from a mask and set the oxygen flowmeter to 0.4-0.8 L/min and the isoflurane vaporizer to 1.5%, then start the surgery.
3. Shave the surgical area with #40 clippers; remove hair from an area 2 times the surgical area, prepare the shaved skin with three antiseptic scrubs of povidone-iodine, followed by 70% ethanol, and cover with a sterile drape.
4. Move the animal under a dissection microscope located in a Biosafety cabinet. Expose the reproductive tract by an incision in the dorsomedial position directly above the ovarian fat pad.
   Note: Critical step: Precise skin incision facilitates wound healing, the use of a scalpel is recommended in step 2.4.
5. Using blunt fine forceps pull the ovarian fat pad carefully through the incision towards the midline, minimizing damage to nerves and major blood vessels.
   Critical Step: If bleeding occurs, stop the surgery and consider transplanting to a different host animal.
6. Under the control of the dissection microscope with help of a 28 gauge beveled needle make a 2-4 mm deep incision into the ovarian fat pad, 3-4 mm above the ovary.
   Critical step: Ensure that the incision only goes through half of the fat pad; puncturing all the way through to the bottom side causes leakage. Be swift and proceed without delay after this step.
7. For cell transplantations, fill a syringe (30 gauge needle) with 10-20 µl of the cell-basement membrane matrix-mixture and inject into the fat pad incision. For uterine tube transplantation, pick up the uterine tube with fine forceps and place in 10 – 20 µl basement membrane matrix kept on ice. Using a 0.1-10/20 µl XL graduated filter tip (shortened by 3 mm) pick up the tissue and basement membrane matrix suspension and release into the fat pad incision.
8. Wait 5 minutes for the basement membrane matrix to solidify and place the reproductive tract back into the peritoneum. Close the muscles with 2 stiches of surgical suture and the skin with two small wound clips or surgical suture.
9. Repeat steps 2.4-2.8 to transplant a PBS-basement membrane matrix-mixture into the contra-lateral fat pad of the animal which will serve as control.
   Critical step: After the surgery place the animal on a heating pad, because heat loss is rapid in anesthetized mice.
10. Let animal recover on a heating pad in the native cage and observe the animal until fully awake from the anesthesia. Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbence. Do not return an animal that has undergone surgery to the company of other animals until fully recovered.
11. Place a handful of pre-wet food pellets inside the cage in addition to dry food on the cage after surgery. Apply post-surgical analgesics if needed for the next few days and examine the wound daily. Apply antibiotics according to the approved animal protocol if wound infection occurs. Remove wound clips 10 days after surgery when the wound has fully closed.

3. Histology and Image Acquisition

1. Graft Collection and Preservation
   1. Prepare 4% paraformaldehyde solution by mixing 26 ml ddH₂O with 4 ml 10x PBS and 10 ml 16% paraformaldehyde solution.
   2. Dissect in one block, the engrafted ovarian fat pad, ovary, uterine tube, 0.5 cm uterus as described in steps 1.1.1-1.1.4. Immediately place in 2 ml 4% paraformaldehyde and fix for 2 hr, at RT. From the same animal, dissect the non-engrafted contralateral ovarian fat pad system transplanted with PBS-basement membrane matrix mixture as a control (see step 2.9). Fix and process in the same way.
   3. After fixation wash tissues three times with PBS for 5 min and incubate overnight in PBS at 4 °C.
   4. For the whole-mount imaging of the ovarian fat pad proceed to step 3.2.1.
      Note: After the image acquisition tissues can be frozen (3.3.1) or processed for paraffin embedding (3.3.2). Overnight incubation in 30% sucrose in PBS at 4 °C improves quality of frozen tissues.
2. Image Acquisition
   1. Place whole-mount ovarian fat pad systems in PBS filled dishes and image using an inverted microscope equipped with epi-fluorescence attachment and high-definition color camera, with 4, 10x objectives, and a stereomicroscope equipped with a fluorescence attachment, color camera and Plan Apo 1xWD70, ED Plan 1.5xWD45 objectives.
3. Histology
   1. Following whole-mount image acquisition (3.2.1) place a 200-500 µl drop of embedding medium for frozen tissue specimens on a 2 x 1 cm piece of laboratory film and using forceps, transfer the tissue to the drop. Pick up the film at one edge and transfer the tissue-medium drop to a 1.8 ml cryogenic vial. Close the vial and plunge into liquid nitrogen. Store frozen tissues at -80 °C.
   2. Alternatively, proceed with the standard paraffin embedding. Use 20-40 volumes of tissue volume for each step.
      1. Immerse tissues once in 65% ethanol for 30 min, shaking gently at RT.
      2. Immerse tissues once in 70% ethanol for 30 min, shaking gently at RT. At this stage samples can be stored for months at RT.
      3. Immerse tissues once in 90% ethanol for 30 min, shaking gently at RT.
      4. Immerse tissues once in 95% ethanol for 30 min, shaking gently at RT.
      5. Immerse tissues twice in absolute ethanol (200 proof) for 30 min, shaking gently at RT.
      6. Immerse tissues twice in chloroform for 30 min, shaking gently at RT.
      7. Immerse tissues once in paraffin for 30 min, shaking gently at 58 °C.
      8. Immerse tissues once in paraffin for 12 hr, shaking gently at 58 °C.
      9. Immerse tissues once in paraffin for 3 hr, at 58 °C.
      10. Place tissues in metal histology molds and fill molds with 58 °C paraffin. Add a plastic histology cassette on top of the paraffin and cool down paraffin to +4 °C. Extract paraffin-tissue block and store at RT.
          Note: Paraffin temperature should not exceed 58 °C for optimal preservation of tissue suitable for immunohistochemical staining.
   3. Alternatively, Prepare for the Paraffin Embedding with Specimen Processing Gel.
      1. Aspirate the PBS and transfer whole-mount tissues to 70% ethanol. Incubate at room temperature for 2 hr. In a water bath, preheat specimen processing gel to 60 °C.
      2. Pipette 200 µl of the specimen processing gel onto a laboratory film-lined Petri dish. Using fine dissection forceps transfer the whole-mount tissue system to the specimen processing gel drop.
      3. Allow the specimen processing gel plug to solidify at room temperature, or solidify the plug for 10 min at 4 °C.
      4. Place the specimen processing gel embedded tissue in histology tissue cassettes sandwiched between biopsy foam pads. Embed in paraffin as in steps 3.3.2-3.3.2.10.
         Note: Specimen processing gel embedding prevents loss and allows preservation of small fragile tissue samples.
         Note: Tissues preserved by freezing or paraffin embedding are suitable for immunohistochemical staining^14,15.