Cell line Xenograft Injections (PJ Keller)

Xenograft Injection Prep ( Click to Download )

Preparing cells for injection:

  1. Determine the total volume of injections (plan on 40 µl per gland plus enough for 1-2 extra glands per cell line/condition).
  2. Thaw out enough matrigel on ice for 50% of the total volume of injections.
  3. Trypsinize cells to a single cells suspension and count.
  4. Put 1 x 106 cells per gland plus enough for 1-2 extra glands per cell line/condition in a conical tube and pellet.
    1. Optional:  wash 1X with PBS or media
  5. Make a solution of 50% Matrigel and 50% media, keep on ice (if cell pellet is going to take up significant volume, you could account for this by making 60% matrigel and 40% media)
  6. Aspirate all media from pellets and resuspend in matrigel/media mix; use enough volume for 30-40 µl per injection.  Try to take into account the approximate volume of the pellet to the best of your ability to estimate when resuspending.
  7. Keep cells in Matrigel:media mix on ice until injection.

Preparation of Anesthetics

Mouse Surgery Protocol ( Click to Download )

Mouse Surgery Protocols
(edited 1/11/96)

A. 2,2,2-tribromoethanol (Avertin™) — From Papaioannou and Fox (1993); ref#2008

  1. Solvents for 2,2,2-tribromoethanol:
    1. Add 2.5 g tribromoethanol (Aldrich) to 5 ml 2-methyl-2-butanol (tertiary amyl alcohol; Aldrich) and dissolve by heating to 50oC with stirring or shaking. Add 200 ml distilled water and continue to stir until butanol is totally dispersed.
    2. Alternatively, ad 1 g tribromoethanol to 50 ml distilled water. Stir for >1 h in foil-covered vessel. 100mg/50ml
  2. Filter-sterilize, then store as 50-100 ml aliquots in dark containers at 4oC.
  3. Warm to 37oC and shake well before using.
  4. Use 0.1 to 0.2 ml/10g body weight administered i.p. – 4mg/10g
  5. NOTES AND CAUTIONS: Avertin is stable for at least 2 years when stored properly. Initial pH of the solution will depend on the pH of the water used to prepare it. Decomposition can result from improper storage. Provided the pH of the original solution was >5, this can be tested by adding 1 drop of Congo Red (0.1%w/v) to 5 ml of anesthetic. Purple color developing at pH <5 indicates decomposition to dibromoacetic aldehyde and hydrobromic acid. If this occurs, these products are toxic and anesthetic should be discarded.

B. Ketamin-Zylazine — From Margaret Delano (Animal Care Office, U. Mass.)

  1. Prepare stock solutions by adding 1 ml of a 100 mg/ml solution of ketamine, 0.5 ml of a 20 mg/ml solution of xylazine and 8.5 ml saline yielding a final volume of 10 ml.
  2. Administer 10 µl/g of body weight for mice 23-32 g.

C. Analgesics for surgery – present federal standards suggest that optimal post-operative care should include analgesics to minimize pain. Bupivacaine can be used as a local anesthesia for surgeries that do not invade the body cavity (e.g. mammary transplants). Buprenorphine is to be used as analgesic for major surgeries in which the body cavity is invaded (e.g. ovariectomy, pituitary transplants, etc.) For references see:(Flecknell et al., 1991). Monitoring for distress includes assessing the attitude, appearance of the fur, membrane color, ability to ambulate for 24 h after surgery. Visual assessment of water and food intake are also included. Body weight measurements can be taken before and after surgery as a quantitative measure.

  1. Bupivacaine – for use as a local anesthetic around sites of surgical incisions of the skin.
    1. Product name: Sensorcaine-MPF (Astra, Inc.) (Bupivacaine Hcl, methyl paraben-free, in sterile isotonic solution) Supplied as 0.25% (2.5 mg/ml) solution
    2. Dose for mice: 5 mg/kg
    3. ml/10g BW: 0.02 ml
    4. Administration: Administer with a fine gauge needle (27 or 30 gauge) on 1 ml syringe. Administer subcutaneously. Inject equal volumes of total 6-8 sites in an ellipse 0.5-1 cm from the planned incision site. Allow 3-5 minutes to take effect. One dosing is sufficient for 6-8 h duration.
  2. Buprenorphine – for use as analgesic for mice
    1. Product name: Buprenorphine (source?); supplied as 0.3 mg/ml solution
    2. Dose for mice: 0.05-0.1 mg/kg
    3. Preparation: 1 ml plus 9 ml sterile diluent
    4. Typical dosing volumes:

      Body Wt (g)

      Dose (mg)

      Volume if undiluted (ml)

      Volume if diluted (ml)


      15 .00075 .0025 .025
      20 .001 .003 .03
      25 .0013 .004 .04
      30 .0015 .005 .05
    5. Administration: Administer pre-operatively or peri-operatively subcutaneously. The duration of action is 8-12 h. Doses can be repeated one time if needed.
  3. Acetaminophen – analgesic for mice
    1. Product name: Acetaminophen, pediatric form 32 mg/ml
    2. Dose for mice: 110-305 mg/kg (suggested 1.5 mg/10 g BW)
    3. Preparation: 3 ml/97 ml water; assumes water consumption of 1.5 ml/10g BW/ 24 h
    4. Administration: Provide diluted in water for 24 h post-surgery

Surgical Procedures

Mouse Surgery Protocol ( Click to Download )

A. Transplantation of mammary epithelium into cleared fat pads

  1. Prepare the following equipment:
    1. Cautery and spare batteries
    2. Surgical tools: DeWecker iris scissors (7mm, sharp-sharp); 2 curved iris forceps; tissue forceps; needle-point forceps; hemostat; sterile sutures
    3. Ear tags
    4. Auto-wound clips and remover too
    5. Avertin on ice
    6. 25 gauge needles and tuberculin syringes
    7. Gloves
    8. Dissecting scope and illuminator
    9. 70% ethanol
    10. Materials for samples: microscope slides, liquid N2, 10% NBF, cryotubes
    11. Petri dish and 1X PBS for tissue transplants
    12. Dissecting boards
  2. Anesthetize mice <12g body weight (3-4 wk old) with 175-250 µl of Avertin.
  3. Make an inverted Y-shaped incision along the ventral throacic-inguinal region to expose the mammary fat pads.
  4. Use the forceps handle to blunt-dissect the skin from the peritoneum. Use pins to hold the internal organs away from the fat pad.
  5. Cauterize the nipple and mammary artery running between the #4 and #5 fat pads and separate the two fat pads.
  6. The mammary fat pad containing the parenchyma is removed using DeWecker iris scissors. The proximal lymph node in the fat pad provides a convenient landmark for the farthest limits of dissection.
  7. Once 2-4 mice recipients are prepared, the transplant tissue can be obtained from donor mice. The tissues should be cut into 1 mm cubes.
  8. Implant the tissue or cultured cells (10 µl of cells at a concentration of 5 x 107/ml) into each fat pad.
  9. Close the skin with four wound clips.

B. Ovariectomy

  1. Shave the back of the mouse.
  2. Make a single mid-line incision along the back.
  3. Lay the mouse on its side and locate one ovary. The ovary is beneath a deposit of white fat that is quite apparent in contrast to the surrounding dark red organs. Make an incision through the peritoneum and pull the ovary out using the sharp jewelers forceps (#6). Note that the kidney can be easily damaged because it is attached to the ovary by loose tissues.
  4. Use the Serrifin clamp to hold the ovary. Place a ligature at the base of the ovary, but try to remain above the Fallopian tubes.
  5. Remove the ovary, then close the peritoneum with one or two stitches.
  6. Repeat the procedure on the contralateral side.
  7. Close the skin with 2-9 mm wound clips.

C. Pituitary transplantation to the kidney capsule

  1. Removal of pituitaries
    1. Prepare sterile PBS on ice for holding the pituitaries.
    2. Cervically dislocate the mice by pinching the neck. This is needed to prevent damage to the brain surrounding the pituitary.
    3. Decapitate the mouse.
    4. Expose the pituitary. This is done by first pulling the skin over the cranium and cutting the muscle overlaying the bone to gain a clear view of the skull. Cut along the sutures on the sides of the skull and remove the bone. Remove the cerebrum and cerebellum with forceps from an anterior to posterior direction to avoid damage to the pituitary.
    5. Outline the perimeter of the pituitary cavity with sharp, half-curved forceps to break tissue attachments. Lift the pituitary gently with minimal squeezing. Both lobes should be apparent under the dissecting scope.
    6. Place the pituitaries in 1X PBS for up to 2 hours.
  2. Implantation into recipients
    1. Equipment
      1. PBS
      2. Suture
      3. Sterile cotton swabs
      4. Trocar
      5. Neosporin opthalmic solution
    2. Make a mid-sagital incision on either side of the mouse. However, the spleen is on the left side.
    3. Make an incision through the peritoneum over the kidney.
    4. Before exteriorizing the kidney, prepare the pituitary implant. With the bevel of the trocar in the proper orientation, draw some saline into the trocar, place the pituitary in the trocar, and draw it completely into the trocar. Dip a cotton swab into PBS.
    5. Use a cotton swab to push the kidney out.
    6. Hold the kidney between the thumb and index finger.
    7. Nick the kidney capsule with sharp forceps. Insert the trocar into the hole in the capsule and well under the capsule. Deposit the pituitary and withdraw carefully the trocar. (If the capsule tears, it will be necessary to try the contralateral kidney or use a different mouse.)
    8. Close the peritoneum with one stitch and add a drop of neosporin solution.
    9. Close the skin with one wound clip.

D. Growth factor implants into mammary fat pad

E. Vasectomizing mice (as described in detail in “Manipulating the Mouse Embryo”) The male mouse is anesthetized with Avertin. The abdomen is shaved and wiped with 70% ethanol. Using sterile instruments, the abdominal skin is cut at a point level with the top of the legs. A similar incision is made in the body wall. Using blunt forceps, the fat pad is pulled from the incision with the attachee testis, epididymus and vas deferens. With a sharp forceps, a hole is poked in the membrane beneath the vas deferens and a loop of silk suture is pulled through. The suture is cut to give two pieces of suture under the vas deferens. Both pieces are tied and a section of the vas deferens between is removed. The procedure is repeated on the other contralateral vas deferens. The testes are placed back into the body cavity and the body wall is closed with two to three stitches. The skin is closed with wound clips. The mice are placed under a heat lamp until recovery which is usually within 20 minutes.

F. Embryo transfers (as described in detail in “Manipulating the Mouse Embryo”) The recipient is anesthetized with Avertin. The lower back is shaved. After wiping the mouse’s back with 70% ethanol, a small transverse (<1 cm) incision is made with a sterile dissecting scissors about 1 cm to the left of the spinal cord at the level of the last rib. The skin is slid around until the incision is over the ovary or fat pad. The body wall is picked up and a small incision is made just over the ovary. The incision is stretched with the scissors to stop any bleeding. The fat pad and ovary are exteriorized and a serafine clamp is attached to the fat pad and laid down over the middle of the back. The infundibulum is located with the aid of a dissecting microscope. A hole is torn in the ovarian bursa and the embryo transfer pipet is inserted into the infundibulum with the aid of forceps. After embryo delivery, the serafine clamp is removed, the ovary is placed back into the body cavity, and the body wall is closed with 1-2 stitches. The skin is closed with wound clips. Mice are placed under a heat lamp until recovery which is usually within 20 minutes.