Osmotic Pumps in Mice and Rats

The usual site for subcutaneous implantation of osmotic pumps in mice and rats is on the back, slightly posterior to the scapulae (shoulder blades). Other regions may be used, provided that the pump does not put pressure on vital organs or impede respiration or mobility. If the pump is implanted subcutaneously without a catheter attachment, the contents of the pump will be delivered into the local subcutaneous space. Absorption of the compound by local capillaries results in systemic administration. With compounds that are absorbed very slowly by the capillaries, a direct vascular connection from the pump may be required.

For subcutaneous pump implantation, perform the following steps:

1. Once the animal is anesthetized, shave and disinfect the skin over the implantation site.
2. Make an incision 1.5 X the diameter of the implant adjacent to the site chosen for pump placement and perpendicular to the long axis of the implant. If the back of the animal is the site of choice, make a mid-scapular incision across the back perpendicular to the spine.
   • Insert a hemostat into the incision, and, by opening and closing the jaws of the hemostat, spread the subcutaneous tissue to create a pocket for the pump. The pocket should be large enough to allow some free movement of the pump (e.g., 1 cm longer than the pump). Avoid making the pocket too large, as this will allow the pump to turn around or slip down on the flank of the animal. The pump should not rest immediately beneath the incision, which could interfere with the healing of the incision.
   • Insert a filled pump into the pocket, delivery portal first. This minimizes interaction between the compound delivered and the healing of the incision. If the pocket is not large enough to hold the implant comfortably, remove the implant and enlarge the pocket as described above.
3. Close the wound with wound clips, sutures or surgical glue. Two clips will normally suffice. In mice, sutures or surgical glue are recommended for comfort.
4. Provide analgesia as indicated in the IACUC protocol.

1. Osmotic pumps can be implanted intraperitoneally in animals with sufficiently large peritoneal cavities Depending on the size of the animal relative to the pump, intraperitoneal implantation can disrupt normal feeding and weight gain for a day or two thereafter. Allow 48 hours for the animal to recover after intraperitoneal implantation before any experimental procedures requiring handling or transport are undertaken.
2. With any substance administered intraperitoneally, whether by injection or by infusion, a majority of the dose may be absorbed via the hepatic portal circulation rather than by the capillaries. For substances which are extensively metabolized by the liver (i.e., have a high “first pass effect”), the intraperitoneal route of administration may produce highly variable concentrations of agent in plasma and consequently highly variable effects. Therefore, the intraperitoneal route should be avoided with agents that have a significant first-pass effect.

For intraperitoneal implantation, perform the following steps:

• • Once the animal is anesthetized, shave and disinfect the skin over the implantation site.
  • Make a midline length-wise skin incision, 1 cm long, in the lower abdomen under the rib cage.
  • Carefully tent up the musculoperitoneal layer to avoid damage to the bowel. Incise the peritoneal wall along the linea alba directly beneath the cutaneous incision.
  • Insert a filled pump, delivery portal first, into the peritoneal cavity.
  • Close the incision in two layers.

Close the musculoperitoneal layer along the linea alba with 4-0 or 5-0 absorbable suture such as Vicryl or PDS in an interrupted or continuous pattern, taking care to avoid perforation of the underlying bowel.

• • Close the skin incision with 2 or 3 wound clips, interrupted sutures or surgical glue. In mice, sutures or surgical glue are recommended for comfort.
  • Provide analgesia as indicated in protocol.

Via a catheter, osmotic pumps can deliver directly into the venous or arterial circulation. Osmotic pumps have been shown to pump successfully against arterial pressure with no alteration in flow.³ The following procedure details placement of a catheter in the external jugular vein; similar procedures can be followed for the carotid artery. In many cases, this site is preferable because of its size and ease of access. Other sites may also be used.

Note: This procedure requires attachment of a catheter to the pump.

For Intravenous Infusion, perform the following steps:

1. Once the animal is anesthetized, shave and disinfect the ventral portion of the animal’s neck.
2. For ease of manipulation during surgery, the animal can be placed in a sterile stockinette and the head and neck exposed for gas anesthesia administration and surgical access.
3. Position the animal in dorsal recumbency and secure its head and anesthetic delivery apparatus in place (if using gas anesthesia).
4. Place a small bolster beneath the animal’s neck to expose the ventral neck more fully.
5. Use a small, sharp scalpel blade to make a single incision from the ramus of one side of the jaw to the tip of the sternum just lateral to the trachea/midline.
6. Gently dissect down through the salivary and lymphoid glands, adipose tissue, and fascia to the external jugular vein, which is superficial to most of the neck musculature. Gently elevate and clean the jugular vein for a distance of 1.5 cm.
7. Tie off the cephalic end of the vein, leaving tails 4-5 inches long.
8. Place two loose ligatures around the cardiac end of the vein. Place hemostats on the cephalic suture and one cardiac suture to provide gentle counter-traction to the vessel.
9. To inhibit vasoconstriction, apply a few drops of 0.5% lidocaine or other vasodilatory substance (at body temperature), and allow time for effect.
10. Use a fine gauge needle (22-20 gauge for rats; smaller (22-25 gauge) for mice) bent at an approximate 90-degree angle to pierce the vessel. Alternately, a small ellipsoidal piece can be cut from the ventral aspect of the vessel with fine iris or micro scissors. Do not cut so much tissue as to weaken the vessel such that it breaks when traction is applied via the rostral ligature ends while passing the cannula.
11. Once the vessel has been pierced, control hemorrhage with gentle traction on the cephalic ligature ends.
12. The free end of the catheter can be inserted into the hole in the vein wall, and advanced gently to the level of the heart (about 2 cm in an adult rat and about 1 cm in an adult mouse). Tie the cardiac ligatures snugly around the catheter, being careful not to crimp the catheter. The cephalic ligature can then be tied around the catheter. Cut the ends of all three ligatures close to the knots.
13. Using a hemostat, tunnel over the neck, creating a pocket on the back of the animal in the midscapular region. Lead the pump into this pocket, allowing the catheter to reach over the neck to the external jugular vein with sufficient slack to permit free head and neck movement.
14. Pass the caudal end of the pump through this tunnel into the pocket.
15. Use a two-layer closure, with one layer of suture in the underlying fascial tissues, and one in the skin. The deep layer should be closed with 4-0 or 5-0 absorbable material such as Vicryl or PDS in a simple continuous or interrupted suture. The skin can be closed with the same material, nonabsorbable suture, stainless steel wound clips, or surgical glue. In mice, sutures or surgical glue are recommended for comfort.
16. Provide analgesia as indicated in the IACUC protocol.
17. Wound clips or ligatures in the skin should be removed in 10-14 days if the animals are to survive longer than 2 weeks.

a. Cranial² Infusion

For cranial infusion, directing the test substance into the bone (calvarium), the following steps, depending on desired implant location, may be followed:

Catheter Placement

1. Once the animal is anesthetized, shave and disinfect the skin over the cranial implantation site.
2. Wash the scalp three times with 1% betadyne solution followed by 70% ethanol.
3. Make a linear skin incision over the intended implant site.
4. Reflect the skin and mucosal tissues laterally to expose the calvarium.
5. Make a separate incision through the periosteal layer, exposing the calvaria.
6. Create one circular craniotomy defect (1 mm) in the bone with sterile burs using a slow speed dental handpiece, under constant irrigation with sterile 0.9% physiological saline solution under aseptic conditions.
7. Co-adapt the periosteal and skin-mucosal flaps, and then suture with interrupted 5-0 nylon, Gore-Tex, or other inert, nonabsorbable sutures.
8. To place the catheter to the cranial infusion site/calvarial defect and link it to the osmotic pump placed subcutaneously see Section E.c.

b. Intracerebral infusion

For intracerebral infusion directing the test substance into the brain follow above steps but make the incision at the intended implant site and drill through the calvarium to access the brain.

c. Subcutaneous Osmotic minipump implantation for either cranial/calvarial or intracerebral delivery of test compound

1. Local application of the test compound will be accomplished by surgical subcutaneous implantation of the osmotic minipump.
2. With the animal still under anesthesia, the subcutaneous implant site is shaved and aseptically prepared.
3. Access to the cranial site is obtained by primary linear incision.
4. Starting from the distal end of this incision, a supra-periosteal dissection is carried out in the retro-scapular area, creating a pouch on the back of the animal.
5. A filled osmotic minipump with attached catheter is inserted into the pouch.
6. The catheter is positioned and glued in place with cyanoacrylate glue, with the catheter aperture being located slightly over the edge of the bone defect or inserted into the cerebrum. .
7. The flap is repositioned and sutured with 5-0 nylon, Gore-Tex, or other inert, nonabsorbable sutures.

In animals surviving after the pump’s active infusion time, the pump must be removed no later than one half-life (t1/2) after the completion of its infusion time. Thus, a pump designed for one week’s infusion must be removed no later than 3 1/2 days after the end of the 7th day or 10 1/2 days after the implant. A pump designed for two weeks’ infusion is removed no later than one week after the end of the two weeks or three weeks after implant.

1. Surgical removal of subcutaneous osmotic pumps is accomplished in the anesthetized animal via a simple skin incision using aseptic techniques.
2. If the pump has been in place longer than a couple of weeks, or the infusate is an irritant, it may be necessary to free the pump from surrounding connective tissue in order to remove it.
3. When the first pump is removed, examine the site. If there is no scar tissue present, the new pump may be placed in the same location. If scar tissue is present, the new pump should be placed at a new site, far enough away from the previous site so as to not interfere with wound healing or with subcutaneous absorption of the infusate.
4. The pump should be removed in the following circumstances:

a. To verify delivery by measuring residual volume

b. To verify stability and bioactivity of the test agent in solution

c. No later than the recommended “explant by” date

d. Longer infusion periods

If approved by the IACUC; to replace it with a fresh pump, in order to infuse for a longer period than the duration of a single pump. (Note that an explanted pump cannot be reused without being decontaminated and sterilized first.)