Oviduct Transfer

The Whittingham (1968) method, which applied a well-described method for the rat (Noyes and Dickman 1961) to the mouse, is the basis of the procedure described below. It is best to practice this procedure first on a cadaver and then on an anesthetized 0.5-dpc pseudopregnant mouse using a dye solution or blue Affigel beads rather than embryos to gain experience in finding the opening of the oviduct (infundibulum). The position of the infundibulum is relatively invariant from mouse to mouse, and, with a little practice, the procedure will become routine.

Materials

  • Mouse embryos (0.5-3.5 dpc)
  • Pseudopregnant female recipient mice at 0.5 dpc

Equipment

  • Fiber optic illuminator (very useful)
  • Forceps, blunt fine with serrated tips
  • Forceps, sharp
  • Forceps, watchmaker’s #5, two pairs
  • Hypodermic needle, 26-gauge ½ inch
  • Lid of 9-cm plastic Petri dish, glass plate, or paper towel
  • Light bulb, 50W
  • Needle, curved surgical (e.g., size 10, triangular, pointed) (optional)
  • Plasticine (optional)
  • Scissors, fine dissection
  • Serrefine clamp or baby Dieffenbach clip (1.5 inch or smaller) (e.g., Roboz Surgical Instrument RS7440; Weiss B950B, or Fine Science Tools 18050-35)
  • Stereomicroscopes (ideally one for the surgery and one for loading the embryo transfer pipette) with transmitted and reflected light
  • Suture, surgical silk (size 5-0) (optional)
  • Syringe, 1-ml
  • Tissues (several rolled into small swabs are useful for soaking up any blood)
  • Transfer pipettes and mouth pipette assembly (see Chapter 4 and Fig. 4.1A,B)
  • Warming plate or heating pad
  • Weight scale
  • Wound clips and applier

Reagents

  • Anesthetic (see Appendix 1)
  • Beads, blue Affigel (BioRad 153-7302)
  • Epinephrine (optional)
  • Ethanol, 70% <!>
  • M2 medium or other HEPES-buffered medium at room temperature

Procedure

  1. Weigh and anesthetize the recipient mouse by IP injection (see Chapter 3).
  2. Place the mouse on the lid of a 9-cm Petri dish, a paper towel, or other supporting material so that it can be easily lifted onto the microscope stage.
  3. Load a transfer pipette with embryos. Because they will be outside the incubator for several minutes, transfer embryos into M2 or other HEPES-buffered medium before loading the transfer pipette.
    1. Take up a small amount of M2 medium into the transfer pipette, then a small air bubble, then M2 medium, and then a second air bubble; repeat until good control is reached with reduction of capillary action (see Fig. 6.1).
    2. Draw up the embryos in a minimal (about 5-7 mm) volume of M2 medium (Fig. 6.1).  Place an additional small air bubble at the tip of the pipette.
    3. Store the transfer pipette (still in the mouth-pipetting device) by pressing the side of pipette into a piece of Plasticine stuck to the base of a stereomicroscope and leave it there until ready to place the embryos in the oviduct. It is also possible to rest the pipette on a tube rack or any other support next to the microscope, making sure that the tip of the pipette does not touch anything.
      BE CAREFUL NOT TO DISTURB THE PIPETTE.
  4. Expose the recipient’s reproductive tract.
    Wipe the back of the mouse with tissues soaked in 70% ethanol. Remove fur if local legislation requires.  Make a small incision in the skin with fine dissection scissors, along the dorsal midline, at the level below the last rib (Fib. 6.3A,B). Wipe the incision with 70% ethanol-soaked tissue to remove any loose hairs.  As discussed above, both oviducts may also be approached from a single incision made at the dorsal midline perpendicular to the vertebral column at the level below the last rib, or from transverse incisions about 1 cm to the sides of the spinal cord.
    Slide the skin around until the incision is over the ovary (orange) or fat pad (white), both of which are visible through the body wall (Fig. 6.3C). then pick up the body wall with watchmaker’s forceps and make a small incision just over the ovary with fine dissection scissors. It is also possible to pierce the body wall using sharp forceps.  Stretch the incision with the scissors (or forceps) to stop any bleeding (Fig. 6.3D). With a curved surgical needle, thread a piece of surgical silk suture through the body wall so that it will be easy to locate later (optional).
  5. Transfer the embryos.
    1. Use blunt fine forceps to pick up the ovarian fat pad and pull out the attached left ovary, oviduct, and uterus (Fig. 6.3E). Clip a Serrefine clamp onto the fat pad and lay it down away from you over the middle of the back, so that the oviduct and ovary remain outside the body cavity (Fig. 6.3F).
    2. Gently pick up the Petri dish with the mouse and place it on the stage of a stereomicroscope with the mouse’s head to the left.
    3. Using the stereomicroscope, find the opening to the oviduct (infundibulum) and the swollen ampulla located underneath the bursa (a thin transparent membrane containing blood vessels that surrounds the oviduct and ovary). Arrange the mouse, oviduct, etc. so that the pipette can enter easily.  For right-handed people, it is most convenient to have the head to the left and the ovary pulled away from you, toward the right side of the mouse and held outside the body cavity with a Serrefine clamp. For left-handed people, reverse the position of the mouse. Place a drop of epinephrine on the bursa to reduce subsequent bleeding, which can obscure the opening of the oviduct and potentially clog the transfer pipette (optional). With two pairs of sharp watchmaker’s forceps, tear a hole in the bursa over the infundibulum. Alternatively, cut the bursa with fine spring scissors. Be careful not to break any large blood vessels. Locate the infundibulum (Fig. 6.3G).
    4. Use blunt fine forceps to cradle the infundibulum gently and then insert the prepared transfer pipette into the oviduct opening (Fig. 6.3H). Blow on the transfer pipette until air bubble or blue beads (if practicing) have entered the ampulla. Air bubbles or blue beads visible inside the oviduct indicate successful transfer.
    5. Unclip the Serrefine clamp and remove the mouse from the stereomicroscope. Use blunt fine forceps to pick up the fat pad and place the uterus, oviduct, and ovary back inside the body cavity.  Sew up the body wall with one or two stitches (optional) and close the skin with wound clips.
  6. Repeat steps 3, 4, and 5 to transfer additional embryos to the right oviduct, if desired.
  7. At the end of the procedure, place the mouse in a clean cage and keep it warm under a 50W light bulb (taking care to cover the eyes) or by placing the cage on a warming plate or heating pad until the mouse recovers from an injected anesthetic. Follow local animal care committee regulations for more details.

Comments

  • It is advisable to check whether ovulation has occurred as expected right after exposing ovaries and before transferring the embryos. Bloody fluid and small clots on ovaries should be readily visible even without a microscope to confirm recent ovulation. Ruptured graafian follicles may be seen under the microscope. Swollen ampulla and naturally ovulated unfertilized oocytes of the recipient surrounded by a cumulus mass should also be visible. If signs of ovulation are questionable, a different recipient should be used if available.
  • An alternative procedure for embryo transfer into ampullae through a tear in the oviduct wall is described by Nakagata (1992). The oviduct wall may be punctured through the bursa using a 30-gauge needle (or acupuncture needle), eliminating the need to open the bursa to access the infundibulum. The fine blood vessels of the oviduct can serve as landmarks for the puncture, which may be difficult to see.