Use of ALZET Pumps in Neonates

ALZET pumps have been used in neonatal animals, such as rats, guinea-pigs, ferrets, pigs and sheep. References on ALZET studies in neonates. This section discusses considerations for using ALZET pumps in neonates, followed by selected references suggesting surgical techniques and general laboratory practices for working with neonates. This is intended as a general overview of some of the suggested methods in the field. Specific questions should be referred to ALZET Technical Support.

A Method for Chronic Drug Delivery in Neonates

Teratology studies aimed at investigating neonatal learning, memory, behavior or development present unique challenges to scientists because they require manipulation of young, very small animals during a critical developmental period. Authors of numerous rodent studies have concluded that environmental manipulations occurring early in life result in physiological and behavioral changes that persist into adulthood. ^1,2,3 Some studies have shown that human handling of newborn rats for as little as 15-20 minutes daily during the first few weeks of life produces neuroendocrine, neurochemical and behavioral alterations in the adult. ^2,4,5 Some methods of drug administration impose severe experimental conditions, such as repetitive handling and injections, which have more dramatic implications for neonatal development. Furthermore, such experimental artifacts are likely to confound and compromise research results.

Effects on Physical and Neurobehavioral Development

ALZET Osmotic Pumps have long been used as an alternative to repeated injections for chronic administration of experimental agents in unrestrained laboratory animals. These small, implantable pumps present an attractive alternative for use in neonates as well. Doucette et al., at the University of Prince Edward Island, demonstrated the value of these miniature infusion pumps as a method for sustained drug delivery in neonatal rats. ⁶ These investigators used 8-day old Sprague-Dawley rat pups that were randomly assigned to one of three treatment groups: ALZET pump implantation, sham surgery, or no surgery. Saline filled ALZET pumps (Model 1003D) were aseptically implanted under the skin of rat pups under isofluorane anesthesia. The pups were allowed to recover from the anesthesia before being returned to their cages. The entire surgical procedure took an average of 10 minutes and was never longer than 20 minutes. Rats in the sham surgery control group received identical treatment except pump insertion. The rats not operated on were left undisturbed. Animals were evaluated at various times over a 72-day period using a standard battery of tests designed to measure physical and neurobehavioral development, such as weight gain, fur development, incisor eruption, startle, visual placing, and others. With the exception of transient decreases in weight gain during the first 24 to 48 hours following pump implantation, no significant differences were found in rats implanted with pumps compared to the control and sham treated rats on any of the parameters evaluated.

Doucette et al. attributed their experimental success, in part, to the careful use of good laboratory procedures to help minimize surgical stress. They emphasized the importance of following strict aseptic technique during the pump implantation in order to decrease the risk of infection. Additionally, they found the use of an inhaled anesthetic, isofluorane, which permits rapid induction and recovery, to be preferable to slower acting inhaled or injectable anesthetics. The authors concluded that the implantation of ALZET pumps, under carefully controlled surgical conditions, does not significantly affect neurobehavioral development in rat pups, thus they represent a viable alternative to repeated injections for sustained drug delivery.

 

Experimental Model of Neonatal Opioid Tolerance and Dependence

Many experimental models of opioid tolerance and dependence use repeated drug administration by bolus injection with a variety of dosing schedules. Such dosing schedules lead to wide fluctuations of opioid concentrations in the central nervous system that may affect the development of tolerance. Additionally, the added stress from repeated handling and injections could affect the development of tolerance as well. Investigators at the Medical College of Virginia have used ALZET pumps successfully to establish an animal model of neonatal opioid tolerance and physical dependence. Using this experimental model, Thornton et al. have characterized tolerance and dependence to fentanyl and morphine in neonatal rats. ^7,8 Their studies indicate that continuous subcutaneous delivery using ALZET pumps is particularly useful since it closely mimics the intravenous route by which opioids are continuously administered to human neonates. ⁷ Another key benefit of the ALZET pumps in these experiments was their ability to maintain stable plasma and tissue opioid levels, thus reducing toxicity associated with widely fluctuating plasma levels typical with conventional dosing methods. Furthermore, the pumps provided a means of chronic opioid delivery that minimized neonatal handling and the stress commonly seen with repeated injections.

Since these initial studies were published in 1997, the Virginia research group has produced a number of publications describing further research on the long-term consequences of opioid tolerance and dependence established during the neonatal stage. ^9.10,11,12 Thornton et al. have now incorporated ALZET pumps as their standard method for chronic opioid administration in neonatal rats. To obtain a complete list of references on the use of ALZET pumps in neonates, a package of information on surgical techniques for neonates, or additional information about ALZET Osmotic Pumps please contact ALZET technical services.

1. Meany MJ, Aitken DH, van Berkel C, Bhatnagar S & Sapolsky RM. Science 1988;239(4841):766-768.
2. Meaney MJ, Mitchell JB, Aitken DH, Bhatnagar S, Bodnoff SR, Iny LJ & Sarrieau A. Psychoneuroendocrinol 1991;16(1-3):85-103.
3. Smythe JW, McCormick CM, Rochford J & Meaney MJ. Physiol Behav 1994;55(5):971-974.
4. Sapolsky RM. Science 1997;277:1620-1621.
5. Liu D, Diorio J, Tannenbaum B, Caldji C, Francis D, Freedman A, Sharma S, Pearson D, Plotsky PM & Meaney MJ. Science 1997;277:1659-1662.
6. Doucette TA, Ryan CL & Tasker RA. Physiol Behav 2000;71:207-212.
7. Thornton SR & Smith FL. J Pharmacol Exp Ther 1997;281:514-521.
8. Thornton SR, Wang AF & Smith FL. Eur J Pharmacol 1997;340:161-167.
9. Thornton SR & Smith FL. Eur J Pharmacol 1998;363:113-119.
10. Choe CH & Smith FL. Pediatr Res 2000; 47(6):727-735.
11. Thornton SR, Lohmann AB, Nicholson RA, & Smith FL. Pharmacol Biochem Behav 2000;65(3):563-570.
12. Lohmann AB & Smith FL. Pediat Res 2001;49(1):50-55.

Selected References: Surgical Techniques & General Lab Practices for Infusion in Neonates

1. Selected References: Surgical Techniques & General Lab Practices for Infusion in Neonates 1. Park, C.M., K.E. Clegg, C.J. Harvey-Clark, and M.J. Hollenburg. 1992 "Improved Techniques for Successful Neonatal Rat Surgery." Lab. Anim. Sci., 41: 508-513.

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Summary of Important Techniques

• Conditioning of pregnant females to handling & key olfactory stimuli
• Housing of animals in uncrowded conditions
• Proper oxygenation of pups during anesthesia
• Careful visual monitoring during anesthesia
• Maintaining pups' body temperatures during surgery & recovery
• Using aseptic technique during surgery
• Sealing incisions with tissue adhesive
• Allowing pups to recover from anesthesia before being reunited with dam
• Tattooing pups instead of tagging them to avoid rejection by dam

Objective

Park et al . required a safe, controllable method of anesthesia that would allow for complicated, lengthy (30 to 45 minutes) eye surgery on day-old rat pups.   They investigated two anesthesia methods intending to surmount the following two primary challenges associated with surgery in neonates:

• Mortality due to anesthesia
• Postoperative mortality due to cannibalism or neglect

Methods

Anesthesia methods evaluated:

• Halothane administered via gas anesthetic machine, permitting precise regulation of anesthetic depth for each animal
• Innovar-Vet, a neuroleptanalgesic drug combination, administered by injection via fine-gauge insulin syringe, and followed by rapid recovery via administration of Narcan, a potent narcotic antagonist

 

Postoperative Care:

• Pregnant females were conditioned for 7-10 days before parturition by slow, gentle stroking of each animal in its cage with vocal reassurance for 5-minute intervals every 4 hours throughout the day
• Pregnant females were housed individually in large cages with a thick layer of bedding
• Pregnant rats were familiarized with the odor of agents used during surgery and postsurgically
• All incisions were carefully cleaned of blood to avoid excessive licking and were sealed with tissue adhesive
• Pups were returned to their cages after surgery until motor activity and the ability to vocalize returned

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Results

Halothane

• Of unoperated pups, 7 of 8 (88%) survived anesthesia; one died of respiratory arrest
• 7 surviving pups appeared healthy and normal when examined days later
• All 63 pups subjected to eye surgery under halothane anesthesia survived
• 7 days later, 97% of remaining pups (those not euthanized) had survived and were apparently normal and healthy

 

Innovar-Vet

• All animals (n=16) treated with Innovar-Vet survived anesthesia
• 7 days later, all pups appeared healthy and normal

2.   Flecknell, P.A., Laboratory Animal Anesthesia: An introduction for research workers and technicians.   1987. London: Academic Press.

• It is essential to maintain body temperature
• Maintain good ventilation and fluid balance
• Use inhalational anesthetics so recovery is rapid and normal feeding is resumed as soon as possible.   (Methoxyflurane is safe and effective.)
• **It is unclear whether induced hypothermia actually produces anesthesia or simply immobility. It's best to use volatile anesthetics until the humaneness of hypothermia is verified.**

3.   Waynforth, H.B. and P.A. Flecknell, Experimental and Surgical Technique in the Rat. 1984. London: Academic Press.

• Before and after surgery or injection, neonatal rats and the hands of the investigator should be rubbed gently with cage bedding material or urine from the mother; this will disguise the smell of the investigator and avoid possible cannibalism
• Alternative possibility: sedate mother before removing young (e.g., with 1 mg/kg i.p. diazepam )

4.   Recognition and Alleviation of Pain and Distress in Laboratory Animals, Committee on Pain and Distress in Laboratory Animals. Washington D.C.: National Academy Press. 1992.

• Consider use of inhaled anesthetic whenever possible, because biotransformation is not required for its elimination and depth of anesthesia can be readily controlled
• Hypothermia may be applicable on neonates that have not yet developed effective thermoregulatory mechanisms; it has a wide margin of safety and appears effective in surgery. It is also useful as adjunct to general anesthesia in cold-blooded animals

NOTE: As mentioned in an earlier reference, the usefulness of induced hypothermia as a viable, humane method of anesthesia has not been firmly established.

5.   Rich, S., C. Grimm, K. Wong, and L. Cesar, 1990. "Gas Anesthesia Setup for Methoxyflurane Use in Small Rodents." 32(1):7.

• Can provide a relatively rapid induction (1-5 minutes)
• Animals can be induced smoothly with no physical restraint, and recovery is quick
• See article for description of exact setup procedure

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General Facts of Methoxyflurane (Metofane)

• Compatible with commonly used preanesthetic and other anesthetic agents
• Should be used with caution in animals with liver disease and toxemia
• Clinical signs are not as well defined as with other inhalation agents
• Pedal and palpebral signs are abolished early, so are not good indicators of the depth of anesthesia
• The depth of anesthesia is judged by the degree of muscle relaxation, the presence or absence of the swallowing reflex, and if the breathing is not controlled, and by the rate and character of breathing
• Causes depression of newborn. To minimize depression, the animal should be induced with the minimal amount of ultra short-acting barbiturate necessary for intubation, should be maintained with methoxyflurane in as light a plane as possible, and surgery should be completed as quickly as possible
• If depression does occur, the administration of oxygen will usually bring prompt recovery

 

Use of ALZET Pumps in Nude Mice

ALZET pumps have been used extensively in nude mice since 1980. References on the use of ALZET pumps in nude mice.

Surgical Considerations for Nude Mice

• Following are selected surgical considerations for working with nude mice.
• All injections and surgery must be performed sterilely in a laminar-air-flow room or under a laminar-air-flow hood by sterilely gowned, masked, and gloved operators and assistants
• "Nude" skin may have less mechanical strength than normal skin, requiring careful suturing with finer a needle and more stitches per unit length
• Anesthesia is the same as for normal mice
• Primary objective of aseptic surgery: to reduce microbial contamination of the incision and exposed tissues to the lowest possible practical level
• Temperature-controlled small water 'blanket' should be placed under the rodent during prolonged surgical procedures to prevent hypothermia
• A cork board, plastic tray, or a few paper towels can be placed under the rodent to minimize heat transfer during short procedures

 

Decontamination of Skin Surrounding Incision Site

• It's important to remove the fur over the incision site, and to decontaminate the skin
• Decontamination of skin should be accomplished without soaking the body of the rodent
• Alternatives for removing fur include: plucking, clipping, shaving, or, in selected instances, depilatories
• After fur is removed, skin must be cleansed and disinfected; begin at the incision site and work outward in circles of increasing diameter

 

Selection and Sanitation of Surgical Table and Associated Equipment

• Rodent surgical area can be a room or part of a room that is easily sanitized and not used for other activities when rodent surgery is in progress
• The area should be subdivided into specific places for cages of rodents awaiting or recovering from surgery, for preparing rodents for surgery, and for performing surgery; this reduces potential contamination by fur, feces, and bedding
• Before surgery, lab bench or table should be cleaned and disinfected; quaternary ammonium disinfectants or 70% alcohol are good choices
• Lab benches in front of open windows, next to doors, or in similar locations where air currents and dust are difficult to control should be avoided
• An area in or in front of an exhaust hood should not be used; a high efficiency particulate absorbent (HEPA)-filtered hood, glove box, or plastic bubble is acceptable

 

Preparation and Sterilization of Surgical Instruments

• Autoclavable tip guards for surgical instruments should be used
• Special instrument trays with rows of soft plastic fingers can prevent damage of delicate instruments
• After use, instruments should be soaked in lukewarm water to remove blood and tissue, washed with free rinsing neutral pH detergent, rinsed thoroughly and air dried
• Use a toothbrush to scrub delicate instruments
• Check the tips of delicate instruments--preferably under a microscope--to make sure they are not damaged or dull, and check grooves to verify that no blood or tissue remains
• Do not use damaged or dull instruments
• Steam or dry heat are preferred methods to sterilize surgical instruments; sterility should be verified through periodic use of biological indicators
• Glass bead sterilizers sterilize unwrapped instruments quickly; cool instruments on sterile surface before use
• Instrument packs sterilized by ethylene oxide must be aerated to remove residual gas
• Some chemical sterilizers can irritate tissue
• Alcohol is neither a sterilant nor a high-level disinfectant

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Maintenance of Sterility Between Rodents

• Contamination can be reduced by segregating surgical instruments according to function
• In repetitive rodent surgery, wiping instruments with 70% alcohol and a sterile swab between rodents can reduce bacterial contamination on a short-term basis
• Use a sterile instrument pack with holders
• Even with the techniques mentioned above, a sterile instrument pack should be used after 4 or 5 individual rodents
• Use clear and lightweight surgical drapes; opaque disposable paper or cloth drapes make it difficult to monitor the respiratory rate of small rodents

 

General Practice Considerations 

• Establish special areas and equipment for use with nude mice only
• Maintenance requires strict microbiological control
• Personnel who enter animal rooms to handle nude mice should be careful not to touch any other animals or equipment; if they must handle other animals, they should start work with nude mice before other daily routine work
• Water and food do not need to be withheld; the inability of mice and rats to vomit prevents regurgitation of stomach content
• An electric light (50-75 W bulb) suspended over one end of the cage is a simple heat source for rodents recovering from anesthesia

Sources:  

"Guide for the Care and Use of the Nude (Thymus-Deficient) Mouse in Biomedical Research", A Report of the Committee on Care and Use of the "Nude" Mouse , Institute of Laboratory Animal Resources Assembly of Life Sciences, National Research Council. Reprinted from ILAR NEWS , Vol XIX (2), 1976.

Cunliffe-Beamer, Terrie L., DVM, MS. "Applying Principles of Aseptic Surgery to Rodents." AWIC Newsletter, April-June 1993, Vol. 4, No. 2.

Nomura, Tatsuji and Naoko Kagiyama. "Importance of Microbiological Control in Using Nude Mice" from the Proceedings of the Third International Workshop of Nude Mice, Vol 1, September 6-9, 1979.

"Anesthesia and Surgery of Laboratory Animals," William J. White, V.M.D., M.S. and Karl J. Field, D.V.M. in Veterinary Clinics of North America: Small Animal Practice, Vol 17(5), 989-1017. [general information on surgical and anesthetic techniques on laboratory animals (including rabbits, guinea pigs, rats and mice)].