Ovarian Fat Pad Transplantation Assay: An In Vivo Technique to Introduce Cells into Ovarian Fat Pad in Mouse Models

Overview

In this video, we demonstrate an assay for transplantation of experimental cells and tissue fragments into the mouse ovarian fat pad. This method can be used to monitor cell regeneration and tumor transformation via non-invasive techniques.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. 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 immunodeficient (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 the recipient mouse with a single intraperitoneal 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 the eyes to prevent dryness while the animal is under anesthesia. Inject the animal subcutaneously with the analgesic Ketoprofen at a 4 mg/ g body weight dosage to treat 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 a site two 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.
   CRITICAL STEP: Precise skin incision facilitates wound healing; the use of a scalpel is recommended in step 1.4.
5. Using fine blunt forceps, pull the ovarian fat pad carefully through the incision towards the midline, minimizing damage to the 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 the 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 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 it into the fat pad incision. For uterine tube transplantation, pick up the uterine tube with fine forceps and place it in a 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 it 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 two stitches of surgical suture and the skin with two small wound clips or surgical sutures.
9. Repeat steps 1.4-1.8 to transplant a PBS-basement membrane matrix mixture into the animal's contra-lateral fat pad, which will serve as a control.
   CRITICAL STEP: After the surgery, place the animal on a heating pad because heat loss is rapid in anesthetized mice.
10. Let the 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 recumbency. 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 ten days after surgery when the wound has fully closed.

Materials

Name	Company	Catalog Number	Comments
28 gauge needle (hypodermic syringe with attached needle)	Kendall	30339	Part Number: 8881500014
Basement membrane matrix (Geltrex)	Invitrogen	A1413202
Ethanol 200 proof	Koptec	V1001	To prepare 70%
Filter tip 0.1-10/20 µl XL	USA Scientific	1120-3810
Isoflurane, 250 ml	Santa Cruz Animal Health	sc-363629Rx
Ketoprofen	Zoetis	4396H
Microscope, stereo	Nikon	SMZ800
Nod/SCID/gamma (NSG) mice	The Jackson Laboratory	5557	NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ
severe combined immunodeficiency (SCID/NCr) mice, BALB/C background)	NCI-Frederick Animal Production Program	01S11