Use of ALZET Pumps in Immunodeficient Mice

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

Surgical Considerations for Immunodeficient Mice

  • Following are selected surgical considerations for working with immunodeficient 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
  • 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

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 immunodeficient mice only
  • Maintenance requires strict microbiological control
  • Personnel who enter animal rooms to handle immunodeficient mice should be careful not to touch any other animals or equipment; if they must handle other animals, they should start work with immunodeficient 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)].

 

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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