IACUC / LARC STANDARD PROCEDURES

PRODUCTION OF GENETICALLY MODIFIED (TRANSGENIC, KNOCK-OUT, KNOCK-IN) MICE

A. Vasectomy

1. Anesthetize a male mouse. Males at least 2 months of age, from any strain with a good breeding performance can be used. B6D2F1 males are a very good strain for this purpose. Weigh the mouse and inject it intraperitoneally with 0.015-0.017 ml of 2.5% Avertin per gram of body weight (see this section below, point 6-F, "Preparation of Avertin").



2. Shave the abdomen. (Shaving can be omitted once expertise is achieved.) Wipe abdomen with 70% ethanol.



3. Open the body wall. Sterilize all instruments by flaming with the alcohol burner. Cut the skin with dissection scissors (a 1.5-cm transverse incision) at a point level with the top of the legs. Make a similar-sized transverse incision in the body wall; put one stitch through the body wall on one side of the incision, and leave a piece of silk suture in place. This helps to find the body wall later. Both testes can be reached through the one incision.



4. Tie off the vas deferens. First locate then pull out the fat pad on the left side using the blunt forceps and a serafine clamp to restrain it. The left testis, vas deferens, and epidydimis will come out with it. The vas deferens lies underneath the testis and can be recognized by a blood vessel running along one side. With a sharp forceps, poke a hole in the membrane beneath the vas deferens and pull through two threads of silk suture, each 5-10 cm long. Tie a double knot with each thread. The knots should be 4-5 mm apart. Sever the vas deferens between the two knots. Pick up the fat pad with the blunt forceps and carefully nurse the testis back inside the body wall.



5. Repeat procedure on the right testis.



6. Sew up the body wall with two to three stitches. Then sew up the skin or clip the skin together with autoclips. (Sutures are recommended because the body clip can affect sexual proclivity).



7. Monitor the mouse hourly until it has completely recovered from the anesthesia. If clear signs of pain, acute discomfort, or adverse reaction to the drug are apparent (e.g., convulsions), the mouse should be euthanized. Following recovery, the mouse should be monitored daily for one week for signs of infection or other persistent problems, in which case the mouse should be euthanized.

REFERENCE: Hogan, B., Costantini, F. & Lacy E. (1986). Manipulating the Mouse Embryo, A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 11724. (Illustrations in this standard procedure are reproduced with permission from this reference.)

	Figure 1

B. Oviduct Transfer

1. Anesthetize the recipient female. First weigh the mouse and then inject it intraperitoneally with 0.015-0.017 ml of 2.5% Avertin per gram of body weight. (The proper dose of Avertin may vary with different preparations, and should be redetermined each time a new stock is prepared.) Place the mouse on a sterile cotton pad so that it can be lifted onto the microscope stage easily.



2. Shave the lower back of the recipient mouse (optional).



3. Load a transfer pipet with embryos. Since they will be outside the incubator for several minutes, transfer the embryos from M16 to M2 medium before loading. Transfer pipets are pulled in advance from BDH hard glass capillary tubes. The narrow part of the pipet should be 2-3 cm in length and 120-180 cm in diameter, i.e., larger than one embryo but smaller than two. The tip should be flush, and should be flame-polished, in order to minimize the damage to the oviduct. The pipet is first filled with light paraffin oil to just past the shoulder. The viscosity of the oil allows one to pick up or blow out the embryos with greater control.

   A small amount of air is taken up, then medium M2, and then a second air bubble, and so on, with up to 6-8 alternating bubbles of air and M2. Next the embryos are drawn up in a minimal volume of medium, then a final air bubble is taken up, followed by a short column of medium. Store the transfer pipet (still in the mouth-pipeting device) by pressing it into a piece of plasticene stuck to the stereomicroscope base plate, and leave it there until you are ready to place the embryos in the oviduct. BE CAREFUL NOT TO DISTURB THE PIPET.

4. Transfer the embryos.



1. Sterilize all instruments by dipping them in 90% ethanol and flaming them with a Bunsen burner. After wiping the mouse's back with 70% ethanol, make a small transverse incision (less than 1 cm) with the dissecting scissors, about 1 cm down from the spinal cord just posterior to the last rib. Wipe the incision with 70% alcohol to remove any loose hairs.



2. Slide the skin around until the incision is over the fat pad or ovary (the fat pad is white and the ovary typically a red sphere), both of which are visible through the body wall. Then pick up the body wall with the watchmaker's forceps and make a small incision just over the ovary. With a surgical needle, thread a piece of silk suture through the body wall so that the body wall will be easy to locate later. With blunt forceps, pick up the fat pad and gently pull out the left ovary, oviduct, and uterus, which will be attached to the fat pad. Clip the serafine onto the fat pad and lay it down over the middle of the back so that the oviduct and ovary remain outside the body wall.



3. Gently pick up the mouse using the cotton pad on which it lies and place it with head to the left on the stage of the stereomicroscope.



4. Under the stereomicroscope locate the opening (infundibulum) to the oviduct and the swollen ampulla underneath the bursa (a transparent membrane over the oviduct and ovary). Arrange the mouse, oviduct, etc., so that the pipet can enter easily. It is most convenient to have the head to the left and the ovary, etc., pulled to the rear with the serafine. With two watchmaker's forceps, tear a hole in the bursa over the infundibulum. Be careful not to tear through any large blood vessels.



5. Pick up an edge of the infundibulum or the bursa near the infundibulum with fine forceps and then insert the pipet down in to the opening to the ampulla. Blow until most of the series of air bubbles behind the embryos have entered the ampulla.



6. Unclip the serafine and remove the mouse from the stereomicroscope on its cotton pad. With the blunt forceps, pick up the fat pad and gently nurse the uterus, oviduct, and ovary back inside the body wall. Sew up the body wall with one or two stitches and close up the skin with wound clips.
   
   
5. Repeat steps 3 and 4 to transfer additional embryos to the right oviduct, if desired.



6. At the end of the operation, return the mouse to its cage and leave undisturbed in a warm, quiet place. It should begin to recover from the anesthetic in about 30-60 minutes.



7. Monitor the mouse hourly until it has completely recovered from the anesthesia. If clear signs of pain, acute discomfort, or adverse reaction to the drug are apparent (e.g., convulsions), the mouse should be euthanized. Following recovery, the mouse should be monitored daily for one week for signs of infection or other persistent problems, in which case the mouse should be euthanized.

REFERENCE: Hogan, B., Constantini, F. & Lacy, E. (1986). Manipulating the Mouse Embryo, A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 11724.

	Figure 2 Embryo transfer pipet showing arrangement of air bubbles, medium, and embryos. Monitoring the position of the air bubbles enables the operator to ensure that all of the embryos have been injected.

C. Uterine Transfer

1. Anesthetize the recipient female mouse. Weigh the mouse and inject it intraperitoneally with 0.015-0.017 ml of 2.5% Avertin per gram of body weight.



2. Shave the lower back of the mouse. This is optional and can be omitted when expertise is achieved. Place the mouse on a small cotton pad, which serves as a stretcher to move the mouse.



3. Expose the uterus, as follows:
   
   
   1. Sterilize all instruments by dipping in 90% ethanol and flaming with a Bunsen burner.
   
   
   
   2. After wiping the mouse's back with 90% ethanol, make a single small longitudinal incision (less than 1 cm) with the dissecting scissors about 1 cm to the left of the spinal cord at the level of the last rib. Wipe the incision with 90% alcohol to remove any loose hairs.
   
   
   

   Figure 3

   
   
   3. Slide the skin to the left or right until the incision is over the fat pad or ovary (the fat pad is white and the ovary typically a red sphere), both of which are visible through the body wall . Then pick up the body wall with a forceps and use a sharp scissors to make a small incision just over the ovary. Insert a piece of surgical silk through the body wall so it will be easy to locate later.
   
   
   
   4. With a blunt forceps, grab the fat pad and gently pull it out along with the ovary, oviduct, and uterus, which will be attached to the fat pad. Clip a serafine onto the fat pad and lay it down over the middle of the back, so the oviduct and ovary remain outside the body wall.
   
   
   
   5. Gently pick up the mouse using the cotton pad underneath it and place it on the stage of the stereomicroscope.
   
   
   
4. Load a transfer pipet with about seven blastocysts. This should result in about five embryos coming to term (75% success rate).




1. Transfer pipets are prepared in advance from a Pasteur pipet or a BDH hard glass capillary by pulling it out to about 200 cm in diameter, and then firepolishing the end. A bend can be placed in the pipet about 1 cm from the end. This allows one to judge how far the pipet has been inserted into the uterus. However, with practice a straight pipet works very well.



2. The pipet is first filled with light paraffin oil up to the shoulder. A small amount of air is taken up (air bubble 1), then M2 medium, and then another air bubble. Next the blastocysts are picked up in a minimal volume of M2 medium (filling about 0.5 cm of the pipet). A third small air bubble and a final portion of M2 at the tip are then taken up in the pipet to hold the group of embryos in place until they are blown to expel them.



3. Store the pipet by pressing it into a piece of plasticene stuck to the stereomicroscope base plate until it is time to place the embryos in the uterus. BE CAREFUL NOT TO DISTURB THE PIPET.



5. Transfer the embryos.



1. Hold the top of the uterus gently with a pair of fine blunt forceps. Use a 26-gauge needle or an intestinal surface needle to make a hole a few millimeters down the uterus. Avoid small blood vessels in the uterine wall. Make sure the needle has entered the lumen and has not become lodged in the wall of the uterus. To test whether the needle has entered the lumen, pull it out slightly. If it slides easily, the needle has penetrated the lumen. Do not move the needle around too much or the wall of the uterus may become lacerated.



2. Keeping an eye on the hole made by the needle, pull out the needle and insert about 5 mm of the transfer pipet. Blow gently until the air bubble closest to the embryos (air bubble 2) is at the tip of the pipet and all of the blastocysts have been expelled.



3. Unclip the serafine and remove the mouse from the microscope stage using the cotton stretcher pad. With the blunt forceps, pick up the fat pad and gently nurse the uterus, oviduct, and ovary back inside the body wall. Sew up the body wall with one or two stitches and close the skin with wound clips.



6. Repeat steps 3, 4, and 5 on the right side if desired. Use a maximum of 8 embryos if transferring into one side only, or a total of 12 embryos per mouse for bilateral transfers.



7. Monitor the mouse hourly until it has completely recovered from the anesthesia. If clear signs of pain, acute discomfort, or adverse reaction to the drug are apparent (e.g., convulsions), the mouse should be euthanized. Following recovery, the mouse should be monitored daily for one week for signs of infection or other persistent problems, in which case the mouse should be euthanized.

REFERENCES:
Hogan, B., Constantini, F. & Lacy E. (1986). Manipulating the Mouse Embryo, A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 11724

Bradley, A. (1987). Production and analysis of chimeric mice. In Teratocarcinoma and Embryonic Stem Cells: A Practical Approach, ed. E.J. Robertson, IRL Press, Oxford, England.

Figure 4 Diagram showing technique of uterine transfer. A 26-guage or sewing needle is used to make a hole all the way into the lumen of the uterus.

Figure 5 After removing the needle, the transfer pipet is inserted into the hole and the blastocysts injected.

D. Anesthesia of Mice

1. Short-term anesthesia (1-5 minutes)

   Anesthesia by inhalation is required for tail biopsy of mice older than 3 weeks, and for collecting blood by intraorbital bleeding.

   Procedure: Inhalation of metafane is recommended instead of ether because of its lower toxicity and clearer dose dependence. Anesthetize the mouse by placing it into a covered chamber containing a cotton swab saturated with metafane until the mouse is unconscious. Observe the respiratory rate, which should decline under anesthesia. Remove the mouse from the chamber immediately if the respiratory rate becomes suddenly rapid.




2. Longer-term anesthesia (10-60 minutes)

   The formulation called Avertin is recommended for longer-term anesthesia during vasectomy of male mice and oviduct or uterine transfers into female mice, although anesthetics such as metafane can also be used.

   Procedure:

   1. Weigh the mouse and inject it intraperitoneally with 0.015-0.017 ml of 2.5% Avertin per gram of body weight. Place the animal back in the cage and wait until it has stopped moving.
   
   
   
   2. Verify anesthesia by confirming a reduced respiratory rate and lack of response to gentle pinching of the footpad.
   
   
   
   3. Perform required surgery within five to ten minutes.
   
   
   
   4. After surgery, the mouse should be placed under a heating lamp to prevent hypothermia. The animal should be observed post-surgically until it returns to consciousness.
   
   
   
   5. Monitor the mouse daily for one week for alertness, movement and feeding. If any adverse effects of the surgery are observed, termination of the experiment by euthanasia (cervical dislocation) is recommended.

E. Preparation of Avertin

A solution of 100% Avertin is prepared by mixing 10 g of tribromoethyl alcohol with 10 ml of tertiary amyl alcohol (Sigma). Dilute 10 ml of this solution to 2.5% in 390 ml isotonic saline (PBS), then sterilize by filtration and aliquot into a series of sterile snap cap tubes. The 2.5% stock solution is stored wrapped in foil at 4^oC. The proper dose of Avertin may vary with different preparations and should be redetermined each time a new 2.5% stock is made by conducting a dose response experiment. Briefly, inject a set of age matched mice with either 0.01, 0.015, 0.017, 0.02, or 0.025 ml/g of the new stock, monitoring completeness of anesthesia and absence of subsequent adverse side effects. The optimal dose typically proves to be around 0.015-0.017 ml/g body weight.

Note on preparation of Avertin: When diluting the alcohol mixture with some commercially available complex phosphate buffered saline solutions, precipitation of the tribromoethyl alcohol may occur. This is due to the presence of calcium and/or magnesium in the PBS. To avoid this, check that the PBS you use is simple sodium phosphate buffered saline (0.8%) and does not contain calcium and/or magnesium, or use the following Tris buffered saline solution:

• 0.8% sodium chloride
• 1mM Tris, pH 7.4
• 0.25 mM EDTA

Note on stability of Avertin: It is the experience of the major transgenic labs at UCSF that 2.5% Avertin which is prepared in PBS, sterile filtered, and stored in the dark at 4^oC is stable for at least one year. However, the tribromoethyl alcohol is unstable as a solid and subject to the generation of toxic degradation products. When a bad stock of this compound is used, the 2.5% Avertin may induce lethality and/or peritoneal damage. If mice do not recover rapidly from the surgery, or appear listless (or die) after several days to a week, then the tribromoethyl alcohol is suspect. In this case, order a fresh bottle and make a new stock of Avertin.