Implantation & Explantation

ALZET osmotic pumps can be implanted subcutaneously or intraperitoneally following the animal size guidelines. ALZET pumps may also be connected to a catheter to deliver the pump contents directly into the venous or arterial systems, the brain, or into any organ or tissue. (See other routes of administration)

Included on our site are guidelines on surgical implantation of ALZET pumps subcutaneously and intraperitoneally, as well as intravenous cannulation and brain infusion.

In addition, DURECT has a video available on CD or download that demonstrates surgical techniques and special applications for the use of ALZET osmotic pumps. This video demonstrates subcutaneous and intraperitoneal implantations, intravenous infusion (via the external jugular vein), localized administration into the central nervous system, gastrointestinal delivery, intrathecal cannulation, and other special applications. The subcutaneous procedure is shown in a mouse and the remainder are performed in rats. This video is available free of charge. Request a copy

Considerations

  • Subcutaneous (SQ) implantation is technically the easiest and least invasive procedure.
  • Volatile inhalation anesthetics are best for most indications in most species as induction and recovery times are shorter and the surgical plane can be maintained for a short or long duration. Injectable anesthetics are an option in some instances.
  • The pumps cannot be left implanted indefinitely. Refer to our guidelines on explantation.

Subcutaneous Implantation

The usual site for subcutaneous implantation of ALZET pumps in mice and rats is on the back, slightly posterior to the scapulae. Other regions may be used, provided that the pump does not put pressure on vital organs or impede respiration. 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 (see Catheter section).

For subcutaneous pump implantation, perform the following steps:

  1. Once the animal is anesthetized, shave and wash the skin over the implantation site.
  2. Make a suitable incision adjacent to the site chosen for pump placement. If the back of the animal is the site of choice, make a mid-scapular incision.
  3. 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.
  4. Insert a filled pump into the pocket, delivery portal first. This minimizes interaction between the compound delivered and the healing of the incision.
  5. Close the wound with wound clips or sutures. Two clips will normally suffice.

Intraperitoneal Implantation

ALZET pumps can be implanted intraperitoneally in animals with sufficiently large peritoneal cavities (see Guide to Animal Sizes). 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 24 to 48 hours for the animal to recover after intraperitoneal implantation.

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 probably be avoided with agents that have a significant first-pass effect.

For intraperitoneal implantation, perform the following steps:

  1. Once the animal is anesthetized, shave and wash the skin over the implantation site.
  2. Make a midline skin incision, 1 cm long, in the lower abdomen under the rib cage.
  3. Carefully tent up the musculoperitoneal layer to avoid damage to the bowel. Incise the peritoneal wall directly beneath the cutaneous incision.
  4. Insert a filled pump, delivery portal first, into the peritoneal cavity.
  5. Close the musculoperitoneal layer with 4.0 absorbable suture in an interrupted or continuous pattern, taking care to avoid perforation of the underlying bowel.
  6. Close the skin incision with 2 or 3 wound clips or interrupted sutures.

Intravenous Infusion (via the External Jugular Vein) in Rats (click here for mice)

Via a catheter, ALZET pumps can deliver directly into the venous or arterial circulation. ALZET pumps have been shown to pump successfully against arterial pressure with no alteration in flow. The following procedure details placement of a catheter in the external jugular vein. 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 (more info)

When cannulating the jugular vein of rats, use the Rat Jugular Catheter (0007710), sold by DURECT Corporation. This catheter fits onto an ALZET Osmotic Pump with no modification and is provided sterile.

 

Step 1. Prepare the pump and catheter (more info).   Note:   In applications involving a catheter, the pump must be primed before implantation (more info).

Step 2. Once the animal is anesthetized, shave and clean the ventral portion of the animal's neck.

Step 3. For ease of manipulation during surgery, the animal can be placed in a sterile stockinette and the head and neck exposed for anesthesia administration and surgical access.

Step 4 . Position the animal in dorsal recumbency and secure its head and anesthetic delivery apparatus in place.

Step 5 . Place a small bolster beneath the animal's neck to expose the ventral neck more fully.

Step 6. 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.

Step 7. 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.

Step 8 . Tie off the cephalic end of the vein, leaving tails 4-5 inches long.

Step 9 . 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.

Step 10 . To inhibit vasoconstriction, apply a few drops of lidocaine or other vasodilatory substance (at body temperature), and allow time for effect.

Step 11 . Use a fine gauge needle (22 - 20 gauge for rats)* 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.

Step 12 . Once the vessel has been pierced, control hemorrhage with gentle traction on the cephalic ligature ends.

Step 13 . 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). 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.

Step 14 . 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.

Step 15 . Pass the caudal end of the pump through this tunnel into the pocket.

Step 16 . 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 in a simple continuous or interrupted stitch, but silk is acceptable for short-term survival studies of 2-4 weeks. The skin can be closed with the same material, nonabsorbable suture, or stainless steel wound clips.* Wound clips or ligatures in the skin should be removed in 1-2 weeks if the animals are to survive longer than 2-4 weeks.

Additional Recommendations for IV Cannulation in Mice

  • When cannulating the jugular vein of mice, use the Mouse Jugular Catheter 0007700), sold by DURECT Corporation. This catheter fits onto an ALZET Osmotic Pump with no modification and is provided sterile.

  • Use a 25-23 gauge needle bent at an approximate 90-degree angle to pierce the vessel.
  • In mice, sutures are recommended for comfort.
  • See a list of references on the use of ALZET pumps for IV delivery in mice.

Sources:

ILAR, NRC (1996) Guide for the care and use of laboratory animals,Washington, D.C.: National Academy Press.

Stepkowski, S. M., Tu, Y., Condon, T. P., Bennett, C. F. (1994) 'Blocking of heart allograft rejection by intercellular adhesion molecule-1 antisense oligonucleotides alone or in combination with other immunosuppressive modalities', Journal of Immunology, 153, 5336-5346.

Tu, Y., Stepkowski, S. M., Chou, T.-C., Kahan, B. D. (1995) 'The synergistic effects of cyclosporine, sirolimus, and brequinar on heart allograft survival in mice', Transplantation, 59(2), 177-183.

Brammer, D. (1999) Personal communication.

Popesko, P., et al. (1992) A color atlas of small laboratory animals, Volume Two, Rat, Mouse & Hamster, London: Wolf Publishing Ltd.

Localized Administration of Agents to the Central Nervous System in Rats

Three kits for performing brain infusions with ALZET pumps are available from DURECT. Click here for detailed info about ALZET Brain Infusion Kits.

Direct access to the CNS via a cannula implanted in the cranium is useful in experimental situations where the test compound has effects on the CNS, but does not cross the blood-brain barrier appreciably. Significant doses can be administered directly to the brain using this technique, which can eliminate the uncertainty of systemic pharmacokinetic variables. Administration usually takes two forms:

  1. Infusion into the cerebrospinal fluid via the cerebral ventricles.
  2. Direct microperfusion of localized regions of solid brain tissue.

Depending on the nature of the compound administered, intraventricular infusion exposes a wide range of brain regions to the infusate. In contrast, direct microperfusion usually results in a very localized exposure in discrete brain structures. The extent to which different compounds are distributed in brain tissue following local infusion is detailed in the following article:

Sendelbeck SL and Urquhart J. Spatial distribution of dopamine, methotrexate, and antipyrine during continuous intracerebral microperfusion. Brain Res 1985, 328:251-258. ( download PDF )

The following instructions are intended for use with ALZET Brain Infusion Kits.

Surgical Procedure:

Tips on adapting for mice

 

Step 1. Anesthetize the rat using either an inhalant anesthetic (such as isoflurane) or injectible anesthetic (such as Xylazine® and Ketamine®, or sodium pentobarbital). Fit the rat into a stereotaxic apparatus.

Step 2. Shave and wash the scalp. Starting slightly behind the eyes, make a midline sagittal incision about 2.5 cm long and expose the skull. With the rounded end of a spatula, lightly scrape the exposed skull area and pat it dry. Scraping should remove the periosteal connective tissue which adheres to the skull, permitting good adhesion of the dental cement which is later used to secure the cannula.

Step 3 . Identify the bone suture junctions bregma and lambda.   With these as reference points, determine and mark the location for cannula placement using the stereotaxic coordinates determined above in Step 1 of the pump preparation section. Drill a hole through the skull at the marked, stereotaxically correct, location. This hole will receive the cannula.

Step 4. Insert the L-shaped cannula, which is attached by tubing to the ALZET pump, through the skull. To facilitate precise placement of the cannula, the tab on the top of the cannula can be attached to the electrode holder of a stereotaxic apparatus. After the cannula is firmly cemented in place, the tab is easily removed with a heated scalpel. Alternatively, this tab may be removed in advance and the cannula placed by hand. After insertion, the cannula's external arm should lie parallel to the surface of the skull with the tubing extending caudally.

Step 5.* Drill a second hole part way through the skull lateral to the cannula. This second hole will be used to receive a small stainless steel screw which acts as an anchor to secure the cannula.

Step 6.* Insert the small anchor screw while taking care not to go entirely through the cranium. Once the screw has been started into the skull, a turn or two is sufficient to secure it. The small anchor screw should extend approximately 1-2 mm above the skull.

Step 7 . Completely dry the skull surface and cover the cannula, the entire implantation site, and the anchoring screw with dental cement. The powdered dental cement can be mixed with its acrylic solvent in a dish and applied. Alternatively, the powder can be placed first and the solvent carefully added to it, taking care to limit both to the implantation site.   Note:   Many researchers use cyanoacrylate adhesive in place of dental cement ( more info ).

Step 8. After the cement has set (about 4 minutes), prepare a subcutaneous pocket in the midscapular area of the back of the rat to receive the osmotic pump. This pocket is created by opening and closing a hemostat to blunt dissect a short subcutaneous tunnel from the scalp incision to the mid-scapular area. The pocket should be large enough to accommodate the pump and permit some pump movement, but not so large as to allow the pump to slip down onto the flank of the animal.

Step 9. Insert the osmotic pump, still attached to the catheter leading to the brain cannula, into the subcutaneous pocket. The osmotic pump should be placed with the delivery port pointing toward the cannula site. When the pump is properly placed, the catheter should have a generous amount of slack to permit free motion of the animal's head and neck.

Step 10. Close the scalp wound with wound clips or interrupted sutures.

Step 11. Remove the animal from the stereotaxic apparatus and place it back into its cage. The animal requires no restraint or handling during the delivery period.

*Note: These steps may be optional when the brain infusion kit 2 or 3 are used.

Verifying Cannula Placement

Upon sacrifice, verify the placement of the cannula and its patency according to the following method. Fix the brain with a suitable fixative (e.g., 4% formaldehyde). Remove the jaw and roof of the mouth of the rat and expose the floor of the brain. Cut the catheter and slowly inject a dye (e.g., Evans Blue) through the catheter toward the cannula. Expose the tip of the cannula and examine the dye stains to confirm its placement. Alternatively, after the cannula is removed, the brain can be fixed, frozen, and sectioned to confirm cannula placement.

Targeted Delivery of Agents to the Central Nervous System (CNS) of Mice:

Infusing agents into the mouse CNS is facilitating new research. The low flow rate and small size of the ALZET Osmotic pump used with the Brain Infusion Kit make an ideal combination for intracerebral delivery in mice.   References on the ICV delivery of agents to mice.

Following are tips on infusion to the mouse brain using the ALZET Brain Infusion Kits:

  • Use the spacers provided with the Brain Infusion Kit, as this will allow proper depth placement of the cannula for the mouse brain.
  • Do not use a stay screw or dental cement as described for a rat brain infusion procedure. The mouse skull is too thin to support a stay screw, and there is not sufficient skin to close the incision over a large amount of dental cement. Preferably, secure the cannula in place using cyanoacrylate adhesive such as Loctite 454 .
  • The upper portion of the plastic cannula which is used for attachment to the stereotax arm should be removed before closing the incision. This part would protrude too far above the mouse skull to allow closure of the scalp incision. It can be most easily removed using a heated scalpel.
  • Proper cranial coordinates for cannula implantation are essential. A new mouse brain atlas by Franklin and Paxinos has recently been published, 1 while two older atlases have been cited with some frequency. 2,3

1 Franklin BJK, Paxinos G; 1997 . The mouse brain in stereotaxic coordinates. Academic Press, San Diego, CA.

2 Sidman RL, Angevine JB, Taber PE; 1971. Atlas of the mouse brain and spinal cord. Harvard University Press, Cambridge, MA.

3 Slotnick BM, Leonard CM; 1975. A stereotaxic atlas of the albino mouse forebrain. Rockville, Maryland; Alcohol, Drug Abuse, and Mental Health Administration.

Explanting ALZET Pumps

Surgical removal of the ALZET pumps is accomplished in the anesthetized animal via a simple skin incision. 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.

The pump should be removed in the following circumstances:

  • To verify delivery by measuring residual volume
  • To verify stability & bioactivity of the test agent in solution
  • No later than the recommended “explant by” date (see table)
  • To replace it with a fresh pump, in order to infuse for a longer period than the duration of a single pump. (Long-term duration references) (Extended duration references)
  • Note that an explanted pump cannot be reused.

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Schedule for Removing Spent ALZET Osmotic Pumps

After its pumping lifetime has ended, an ALZET osmotic pump becomes an inert object for a period of time lasting about half again as long as the pump’s specified pumping duration. After that time, because of the continued osmotic attraction of water into the pump, it may swell and begin to leak a concentrated salt solution, resulting in local irritation of tissues around the pump. Therefore, DURECT advises explanting spent ALZET osmotic pumps according to the following schedule:

 

ALZET Pump Model No.
Explant Pump By*
1003D
Day 4.5
1007D
Day 10.5
1002
Day 21
1004
Day 42
2001D
Day 1.5
2001
Day 10.5
2002
Day 21
2004
Day 42
2006
Day 63
2ML1
Day 10.5
2ML2
Day 21
2ML4
Day 42

 

*Data shown correspond to the nominal duration. Actual explant date should be calculated using the exact specifications for each lot of pumps.

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Pump Advantages

  • Ensure around-the-clock exposure to test agents at predictable levels
  • Permit continuous administration of short half-life proteins and peptides
  • Provide a convenient method for the chronic dosing of laboratory animals
  • Minimize unwanted experimental variables and ensure reproducible, consistent results
  • Eliminate the need for nighttime or weekend dosing
  • Reduce handling and stress to laboratory animals
  • Small enough for use in mice or very young rats
  • Allow for targeted delivery of agents to virtually any tissue
  • Cost-effective research tool

 

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Researchers are saying...

“Galantamine was administered via…osmotic pump… A minipump was used in order to better mimic the daily multiple dosing regimen given to patients with Alzheimer’s disease.” Wenk et al., European Journal of Pharmacology 2003;453:319-324.