IACUC Approval Form Guide
www.alzet.com
2008
ALZET Osmotic Pump Implantation (IACUC Approval Form Guide):
A guide for filling out the Institutional Animal Care & Use Committee protocol approval
form using ALZET® Osmotic Pumps.
Disclaimer
About the Pumps
Guide Description
Responsibility
Literature Search
Justification for Pump Use
Pre-Surgical
Anesthesia Induction
Injectable
Volatile
Volatile Induction
Surgical Preparation
Surgical Area
Subcutaneous Implantation
Intraperitoneal Implantation
IV
IV (mice)
CNS
CNS (mice)
Post Operative Care/Analgesics
Pump Specifications
Bibliography
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Disclaimer
The information listed in this document is based on published research articles and consultations
with experienced veterinarians. This information is intended to serve as a resource when filling out
an IACUC protocol approval form for the use of ALZET Osmotic Pumps in your study. This
document should be considered a supplemental resource to your own institution’s policies,
guidelines, protocols, and suggestions. The information below is not “all inclusive” and may
require additional information to be provided by the researcher.
Title of Procedure
Surgical Implantation of ALZET Osmotic Pumps in Rodents
About the Pumps
ALZET® Osmotic Pumps are miniature, infusion pumps for the continuous dosing of laboratory
animals as small as mice and young rats. These minipumps provide researchers with a convenient
and reliable method for controlled agent delivery in vivo, while avoiding the need to handle the
animal during the dosing period. The ALZET pumps allow for continuous delivery from one day to
six weeks with a single pump, without the need for batteries or pre-programming.
Description of the procedures covered by this IACUC guide
This guide should help standardize the procedure by which an osmotic pump system, capable of
controlled rate delivery of compounds, drugs and or agents, may be implanted in murine models.
This document is intended to serve as a resource when preparing an IACUC approval form.
Information may be copied and pasted into your documents as needed. If you need help choosing
a pump, see our comprehensive checklist for use here: Checklist
Responsibility
The user should follow manufacturer’s guidelines for pump selection appropriate to animal size and
desired application. In addition, precautions for agent/vehicle compatibility with delivery device(s)
must be considered and or tested.
The user should adhere to aseptic principles when preparing the agent/vehicle and during filling
and priming of delivery device(s).
The user should follow current guidelines for veterinary anesthesia and aseptic surgical
procedures, as adapted to common laboratory rodents, when performing implantations.
Literature Search
If references are required to help support your study, you can find over 10,000 citations available
on our website under the “Research Applications” section or you can request a custom search
here: Bibliography Request
For citation purposes, please use the following format:
ALZET® Osmotic Pumps
Durect Corporation
PO Box 530 Cupertino, CA 95015
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Justification for Use of ALZET Pumps
ALZET pumps are commonly used as a humane alternative to dosing methods associated with
higher stress levels, such as repeated injections or tethered infusion systems. Animal stress caused
by these dosing methods can obscure research data and affect study reproducibility. ALZET pumps
enable scientists to refine their animal treatment by eliminating the added stress of external
connections and repetitive animal handling. Since ALZET pumps continuously deliver a precise
dose, they ensure that all animals are properly dosed during the study period, thus allowing
scientists to reduce the number of study animals needed to achieve statistical significance. In
addition, the ALZET pumps can be used for in vitro studies where animals need to be replaced
and controlled delivery is still required.
Humane Benefits
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No external connections necessary for infusion
Risk of infection is minimized (compared to externalized infusion systems)
Minimizes animal handling and stress associated with other repetitive dosing methods (i.e.,
injections, gavage, tethered infusion systems)
Animals are unrestrained and able to move freely
Obviates need for frequent animal handling, which may interfere with animal behavior and
other critical study parameters
Animals may be group housed
Eliminates the possibility of a missed or mistimed dose
Eliminates over dosing and under dosing caused by repeated injections (reduces variability
of drug levels in blood and tissues)
No pre-programming, batteries, or complex software required
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Pre-surgical Considerations
Healthy animals from a known source and microbiological status should be acclimatized to the
laboratory environment for a minimum of 72 hours.
Perform a pre-surgical assessment of:
1) Activity
2) Skin/coat characteristics/integrity especially at implantation site
3) Occulo-nasal-oral areas moist with clear fluid, yet free of mucopurulent or other discharge,
swelling or injury
4) Respiratory rate and effort barely discernable
5) Individual or group body weights
Withhold dry food, but not water for 4-12 hours before surgery. This reduces weight of ingesta in
oropharynx and GI tract such that respiratory efforts and venous return to the heart are not
impaired when animals are placed in recumbency under anesthesia.
May give moist or gel diets or enriched water (make sure animals accept additives to water 48
hours prior to removing food)5
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Anesthesia Induction (General)
Animals and occupied animal cages (including induction chambers) should be placed on preheated
circulating water heating pads set to 90-100F. This is to prevent hypothermia, slowed metabolism,
prolonged induction, and recovery times.
A “Balanced Anesthetic Regiment” is recommended, which includes pre-emptive analgesia1
administration with Buprenorphine or Butorphanol delivered SQ 15-60 minutes before induction.
This can easily be administered to the entire group of animals by drawing up enough to dose all
with a single syringe, based on heaviest body weight (it is recommended to dilute with sterile saline
so that the injection volume for each animal is between 0.1 and 0.2 ml). Attach a butterfly
extension with small gauge needle (22-25 x 5/8 inch) so that the animals can be dosed SQ into
tented scapular skin with minimal restraint in their home or transport cages. Change butterfly
between cages/groups or as needed. The long duration of action (4-12 hours) of these agents
allows them to: 1) act as a pre-anesthetic, to facilitate further handing and decrease anxiety/stress,
2) decrease the amount of anesthetic agent(s) to maintain surgical plane 3) alleviate any potential
pain during the immediate post operative period. You may also want to look into NSAID’s 1 (nonsteroidal anti-inflammatory drugs) as they are gaining popularity due to their long acting effects.
Parenteral administration of (100 F or 40 C) sterile isotonic fluids, such as physiological saline or
LRS, should be given at a rate of 1-3% of body weight (SQ or IP), depending on the intended site
of pump placement. These should be administered upon anesthetic induction so they can be
absorbed to offset hypotonic effects of general anesthetics.
Injectable Anesthetics1
Induce surgical plane with an IP injection of Ketamine and Xylazine or some other alpha-adrenergic
agonist such as “Domitor.” Do not inject IM even if planning to implant IP as most of the agent will
track back out of the needle path and end up SQ, which results in delayed induction and failure to
reach a surgical plane. If desired, the alpha-adrenergic component can be reversed with an
antagonist. The effects of the Ketamine may be diminished, but not reversed with the respiratory
stimulant doxapram.
Recommended drawing of anesthetic is as follows:
Per individual body weight or based on lowest body weight in-group X 2. Thus, have twice as much
as is needed. Administer half in a single IP bolus using good head-down restraint into lower lateral
abdominal quadrant to reduce risk of intraluminal injection into GI.
Place animal in clean paper towel lined cage so particulate bedding will not obscure airways or
damage corneas while going under the influence. Once the animal has lost its righting reflex
proceed with surgical preparation of implantation site(s). If animal does not loose it’s righting reflex
within 5-10 minutes, either remove from study (as anesthetic may have been injected
intraluminally into GI tract), or give an additional dose.
Volatile Anesthetics1
Most common is isoflurane, but halothane is acceptable, though poses more of a health risk to
personnel. The most common carrier gas is 100% medical grade oxygen, but an oxygen enriched
atmospheric mixture or one containing nitrous oxide may also be used. Preemptive analgesia (as
described above) is also recommended, as it will decrease the amount of volatile anesthetic
required.
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Induce in a suitable chamber, which is transparent, air/water tight, and free of sharp/rough
surfaces. The chamber should be large enough to accommodate several animals without crowding
and easily enable retrieval of anesthetized animals, but not so large as to be wasteful of oxygen
and anesthetic agent.
Ideally the chamber should be in a fume or externally vented hood/area, but can also be attached
to an active or passive scavenging system through a charcoal canister. The canister should be
changed regularly.
The chamber should be attached via air and watertight tubing to a calibrated precision vaporizer.
Many commercial units are available or can be adapted from surplus human or larger animal units.
Alternatively, separate systems can be used for induction and maintenance during prep and
surgery. Always leak check systems with carrier gas before each procedure-day. In addition, check
carrier gas pressure and anesthetic reservoir level since changing/filling during procedure increases
potential operator exposure. Always keep at least one syringe of injectable anesthetic for
emergency use if volatile system fails during procedure.
Volatile Induction1
Place animal(s) into chamber, secure lid, and fill with anesthetic gas at 2-4% with a flow of 1-2
L/min. If animals are immature or in a disease state, you may want to preoxygenate them for a
few minutes before adding the anesthetic. Once animals begin to show signs of staggering you
can reduce the flow rate, but not the percent of anesthetic.
Observe animals constantly until loss of righting reflex, and then place them on a snuggly fitting
nose cone. Make sure the cone fits securely over muzzle, but does not abrade corneas. Keep
animals positioned such that the neck is extended to avoid tracheal narrowing. Remember not to
prolong or put undue pressure on the thorax, especially when animal is in ventral or lateral
recumbencey, as it can inhibit respiration and lead to hypoxia and or death.
If attempting to intubate or perform a quick surgical preparation without anesthetic maintenance,
(not recommended unless animals are already devoid of hair) leave in chamber several minutes
longer until significant change in rate and depth of respiratory movements are noticed. Then
perform intubation with appropriate diameter and length tube. Intratracheal intubation in rodents is
possible but requires precision and practice, hence will not be covered here. Methods can be found
in published in literature1.
Surgical Preparation
Place animal on pre-warmed circulating water heating pad, covered with absorbent material to
reduce hypothermia and decrease metabolic rate.
1. Immediately apply ocular lubricant since antiseptic solutions and loose hair can abrade
corneas. Administer fluids as described above to counter act hypotension, caused by
anesthetic agents. This will also help with vessel dilation if cannulating for IV delivery.
2. Shave hair with clean, well-lubricated fine blade clippers. Be sure to periodically check the
clippers as the heat generated can cause thermal burns thus delaying the healing process.
If many animals are being shaved, it is recommended to have multiple clippers so each unit
can rest, clean, and lubricate, while using the other. Clip at least a 1-2 cm border around
implantation site(s). For SQ implantation, clip both the insertion and seating areas so you
can assess healing and pump implantation site(s). It is ideal to have the pump seated
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away from insertion site so there is no pressure on the healing incision. Placing a gentle
two-way traction on skin facilitates clipping. It is best to clip in a designated area slightly
aside from where you will prep skin, so loose hair is contained. Remove loose hair with dry
or just damp cloth or with vacuum suction or compressed air or hair dryer on low or no
heat.
3. Shaved skin should be cleaned of any visible contamination with a dilute antiseptic soap or
solution. Avoid excessive water or alcohol as evaporation facilitates hypothermia. In
rodents that are not visibly soiled it is recommend to simply wipe with antiseptic solution 23 times using freshly opened gauze, wipes, or cotton tipped applicators. It is important to
assure at least 3-5 minutes of contact time with skin before surgical incision is made and to
wipe in a single pass outward motion from incision site towards hair. It is suggested to
avoid detergents as they tend to irritate skin and may lead to scratching during the healing
process. Avoid iodine-based products especially in nude rodents as they can cause
irritation and discolor skin, such that postoperative observation is more difficult. Assess
anesthetic depth by pinching interdigital skin, ear tip, or lip margin with fine rat toothed
forceps. This mimics the full thickness penetration caused by a surgical incision, in a smalldefined, highly innervated site. If an animal shows a withdrawal reflex (twitches in the
pinched area) more anesthetic is needed; wait at least 5 minutes before proceeding. Move
animal to surgical area.
Surgical Area
Animal should be placed on a clean, dry, absorbent, pad, covering a circulating water heating pad
set at approximately 90 – 100F. Two or three sets of autoclaved instruments and trays should be
prepared. The trays should contain the following:
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A tray of high-level disinfectant and a tray of sterile physiological saline.
Optional drying tray. This way one set can be immersed in the high level
disinfectant while the other is being used and then rinsed and dried before using
again. This allows sufficient contact time with the high level disinfectant between
uses. A glass bead sterilizer can also be used, but you must be cautious to avoid
overheating the instruments (have a dipping container of sterile saline or water
and use another cold instrument to transfer from glass beads to cooling rinse).
Draping rodents can be problematic especially when planning to perform two incisions at different
sites such as with an intraperitoneal catheter connected to a SQ pump. Commercially prepackaged
disposable drapes are available, which can be cut down to size such that one can serve a few
animals in a single session. A traditional 4-corner drape is impractical for rodents and with a little
practice the right dimension can be cut for each procedure. A version of the 4-corner drape can be
done with unfolded sterile gauze pads placed over 1-4 sides of the site. Another option, especially
when planning two distinct incisions, is to use commercially available adhesive drape or a sterile
stockinet. A commonly used option is to drape one area at a time and perform an abbreviated
prep with antiseptic on the second site with spray bottle or cotton tipped applicator swab.
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1. Subcutaneous Implantation Procedure
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
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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.
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. The clips
or sutures can be removed 7-10 days post procedure.
6. An analgesic1 should be given post-operatively as needed.
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2. Intraperitoneal Implantation Procedure
ALZET 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 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 that 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 sutures 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. The clips or
sutures can be removed 7-10 days post procedure.
7. An analgesic1 should be given post-operatively as needed.
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3. Intravenous Infusion (via the External Jugular Vein) in Rats
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)
Prepare the pump and catheter (more info). Note: In applications involving a catheter,
the pump must be primed before implantation (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.
1. Once the animal is anesthetized, shave and clean 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 anesthesia administration and
surgical access.
3. Position the animal in dorsal recumbency and secure its head and anesthetic
delivery apparatus in place.
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 countertraction to the vessel.
9. To inhibit vasoconstriction, apply a few drops of lidocaine or other vasodilatory
substance (at body temperature), and allow time for effect.
10. Use a fine gauge needle (22 - 20 gauge for rats)* bent at an approximate 90degree 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.
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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). 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 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 within 1-2 weeks if
the animals are to survive longer than 2-4 weeks.
Additional Recommendations for IV Cannulation in Mice
See cannulation techniques for rat. When cannulating the jugular vein of mice, use one of the
Mouse Jugular Catheters (0007700, 0007701, or 0007702), sold by DURECT Corporation. Any of
these catheters will fit onto an ALZET Osmotic Pump with no modification and they are provided
sterile.
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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 in mice.
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4. CNS Infusion (via Brain Cannulation) in Rats
Direct access to the CNS via an ALZET Brain Infusion 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.
1. Anesthetize the rat using either an inhalant anesthetic 1 (such as isoflurane) or
injectable anesthetic (such as Xylazine® and Ketamine®, or sodium
pentobarbital). Fit the rat into a stereotaxic apparatus.
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.
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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 apparatus. Drill a hole through the skull at the marked,
stereotaxically correct, location. This hole will receive the cannula.
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.
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.
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 12 mm above the skull.
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).
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.
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.
10. Close the scalp wound with wound clips or interrupted sutures.
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 kits 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
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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.
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5. CNS Infusion in 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 ICV delivery in mice.
Following are tips on infusion to the mouse brain using the ALZET Brain Infusion Kits:
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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 stereotaxic
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 mouse brain atlas by
Franklin and Paxinos is a popular choice, 4 while two older atlases have been cited with
some frequency. 2,3
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Post-operative Care/Analgesics (General) 1
Post-operative care should be an extension of proper anesthesia. Provided they do not interfere
with the research focus, analgesics and proper post-operative care are strongly suggested when
performing surgeries, including the implantation of ALZET Osmotic Pumps. See the following list of
references where researchers using the ALZET Osmotic Pumps, cite their use of post operative
care techniques. The following should be considered when providing post-operative care:
Recovery room environment
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The room should be warm and quiet. Lighting should be low, but sufficient enough to
allow proper examination.
The temperature should be 27-30 Celsius for adult animals and 35-37 Celsius for neonates.
For adults this can be reduced to 25 Celsius once the animal has recovered from the
anesthetic.
Caging and Bedding
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Animals should be allowed to recover in their normal cages, a recovery room, or an
incubator.
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Do not allow small rodents or rabbits to recover from anesthesia in cages with sawdust or
wood shavings for bedding (it will stick to eyes, nose, and mouth). Instead, use synthetic
bedding.
Due to cough and swallow reflexes being suppressed during recovery, attempt to minimize
risk of airway obstruction.
Respiratory Depression
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Anesthesia agents can produce respiratory depression and often times this depression is
extended into the post-operative recovery period. In addition, this depression can increase
post-operatively. Hence it is important to monitor the respiratory system in order to
prevent severe hypercapnia and hypoxia.
Use a monitoring device such as a pulse oximeter or a thermistor if the former is
unavailable.
Observe the animal regularly and record the respiratory rates.
If depression is observed, it should be countered with a respiratory stimulant (e.g.,
doxapram and by the administration of oxygen). Since doxapram has a short duration of
action (10-15 minutes), it may be needed in repeated doses or by a continuous infusion.
Fluid Therapy
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Voluntary water intake of all animals should be recorded post-operatively.
Dehydration can compromise the recovery of the animal and should be avoided by
administering fluids post-operatively.
Fluid requirements for most animals are 40-80 ml kg-1 every 24 hours (vomiting, diarrhea,
and other abnormal losses may increase this need).
If animal is conscious, the fluid is best given orally.
If animal is unable/unwilling to accept the fluid, then dextrose-saline (4% dextrose, 0.18%
saline) or saline (0.9%) can be given subcutaneously or intraperitoneally. See table
below:
Mouse (30 g)
Rat (200 g)
Subcutaneous (ml)
1-2
5
Intraperitoneal (ml)
2
5
Infection Prevention
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
The ALZET Osmotic Pumps, ALZET catheters, and other ancillary products sold by DURECT
Corporation are provided sterile. Care should always be taken to use aseptic surgical
techniques and maintain the sterility of the products being used. Doing so may prevent
the need for routine, post-operative, antibiotic administration.
Since animals may soil their wounds with feces and urine, the administration of
prophylactic antibiotics may be useful in minimizing the risk of infection.
For a list of antibiotics and suggested doses for each species see the following:
(Table 6.2)1.
Pain Relief/Analgesics

Proper pain assessment is integral in minimizing post-operative pain. Some key areas to
observe for pain assessment are the following: activity, appearance, temperament,
vocalizations, feeding behavior, and alterations in physiological variables.
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IACUC Approval Form Guide

www.alzet.com
2008
Analgesics can be divided into two groups: opioids or narcotic analgesics and non-steroidal
anti-inflammatory drugs (NSAIDs) like aspirin. For a complete table of analgesic doses see
the citation at the end of this document (Table 6.3) 1
Pump Specifications
ALZET Pump Model
1003D, 1007D, 1002,
1004
Reservoir Volume
100 µl
Complete Osmotic Pump
Model Numbers
2001D, 2001, 2002,
2004, 2006
200 µl
2ML1, 2ML2, 2ML4
2 ml
Length (cm)
1.5
3.0
5.1
Diameter (cm)
0.6
0.7
1.4
Weight (g)
0.4
1.1
5.1
Total Displaced Volume
(ml)
0.5
1.0
6.5
Pump Body Materials
Outer Membrane
Drug Reservoir
Cellulose Ester Blend
Thermoplastic Hydrocarbon Elastomer
Note: All items are supplied sterile. Pumps cannot be reused.
Bibliography
Flecknell P.A. Laboratory Animal Anaesthesia, second edition; A practical introduction for research
workers and technicians. Braintree Scientific
1
Sidman RL, Angevine JB, Taber PE; 1971. Atlas of the mouse brain and spinal cord. Harvard
University Press, Cambridge, MA.
2
Slotnick BM, Leonard CM; 1975. A stereotaxic atlas of the albino mouse forebrain. Rockville,
Maryland; Alcohol, Drug Abuse, and Mental Health Administration.
3
Franklin BJK, Paxinos G; 1997. The mouse brain in stereotaxic coordinates. Academic Press, San
Diego, CA.
4
The Biology and Medicine of Rabbits and Rodents. J. E. Harkness and J. E. Wagner. 1989
5
For more information regarding the ALZET Osmotic Pumps, or to request a complimentary surgical
procedures video, please contact ALZET Technical Support at: 1-800-692-2990 or
alzet@durect.com
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