Substance Administration
Administration by Injection
The injection of substances directly into the body requires strict asepsis to avoid complications. Injected substances and the needles/syringes used to inject substances must be sterile. To prevent contamination of multi-use vials, the rubber top of the injection vial should be swabbed with alcohol prior to use and a new sterile needle must be used to access the vial. Potential complications for all routes of injection include infection, local irritation, pain and damage to surrounding tissue. Video demonstrations of various injection routes are available on the RAT Procedures With Care website. A video demonstration of the retro-orbital injection route is available on the ARP website. Listed below are recommended injection volumes for specific injection routes. Deviations from recommended injection volumes and routes require prior IACUC approval. Please see the references listed at the bottom of the page for more information.
Factors to be considered when selecting an injection route include:
- Pharmacology of the substance administered
- Species and size of animal used
- Final effect desired (e.g., local or systemic)
- Minimization of stress and discomfort to the animal
Intraperitoneal
Potential complications:
- Inadvertent injection into various abdominal organs and subcutaneous, retroperitoneal or intramuscular spaces
- The injection should be made into the lower right abdominal quadrant of the mouse or rat to avoid hitting the liver and intestinal organs of the upper quadrants, the bladder on the lower midline and the cecum in the lower left abdominal quadrant.
- Chemical peritonitis
- Injected material should be sterile, isotonic and nonirritating
- Injection of large volumes can lead to pain, peritonitis, formation of fibrous tissue, perforation of abdominal organs, hemorrhage and respiratory distress
- Tissue damage within the abdominal cavity
- Repeated IP administration can result in a cumulative irritant effect and needle-induced damage of the peritoneum
- Poor injection technique can lead to laceration of abdominal organs or blood vessels
- Hypothermia and distress if large volumes of cold substances are administered
Volumes for intraperitoneal injection:
Mouse = 20 ml/kg (maximum: 40 ml/kg) body weight; do not exceed this recommendation or a total volume of 1.0 ml per mouse without prior IACUC approval. (See references below for additional information. Published dose ranges vary greatly and acceptable dosages are highly dependent on animal size and the experimental situation.)
Rat = 10 ml/kg (maximum: 20 ml/kg) body weight; do not exceed this recommendation without prior IACUC approval. (See references below for additional information. Published dose ranges vary greatly and acceptable dosages are highly dependent on animal size and the experimental situation.)
Hamster = 3-4 ml total volume.
Intravenous
Potential complications:
- Bacteremia/septicemia
- Extravascular delivery of substance administered leading to local soft tissue damage, infection, pain and/or tissue death
- Vascular occlusion, emboli and thrombosis if substances containing particulate material or low pH precipitate in the bloodstream
- Hemolysis, coagulation or anaphylaxis depending on the substance injected
Maximum volumes for intravenous injection:
Mouse: Tail vein and retro-orbital = 0.2 ml
Rat: Maximum volume (bolus) = 5 ml/kg
Hamster: Maximum volume (bolus) = 0.3 ml
Video demonstration:
Intravenous tail vein injection in the mouse.
Intramuscular
Potential complications:
- Pain
- Muscle damage and/or death of tissue
- Irritation or damage to nearby nerves
- Generally not useful in small rodents due to small volume that can be safely administered without tissue damage
Maximum volumes for intramuscular injection:
Mouse = 0.05 ml/site; maximum of 2 sites
Rat = 0.2 ml/site; maximum of 2 sites
Hamster = 0.05 ml/site; maximum of 2 sites
Intranasal
Potential complications:
- Aspiration pneumonia
- Suffocation
- Inaccurate dosing due to sneezing (deep sedation or light anesthesia may decrease this)
Maximum volumes for intranasal administration:
Mouse, rat and hamster = 50 microliters
Subcutaneous
Non-irritating substances can be administered subcutaneously in almost any area of the body where the skin overlying the site is loose enough to allow for volume expansion. Typical sites include the flanks and dorsal shoulder regions. See references below for additional information. Published dose ranges vary greatly and acceptable dosages are highly dependent on animal size and the experimental situation.
Volumes for subcutaneous administration:
Mouse = 10 ml/kg (Maximum possible: 40 ml/kg), not >1.0 ml per site
Rat = 5 ml/kg (Maximum possible: 10 ml/kg total), not >5.0 ml per site
Hamster = 3-4 ml total
Video demonstration:
Subcutaneous injection in the mouse
Dehydration and Fluid Therapy in Rodents
Hydration refers to the amount of water in the body. Adequate hydration is required to maintain normal body function. Dehydration occurs when water loss increases or water intake decreases to the extent that total body water falls below normal. Dehydration is a common complication of illness or injury and is likely to occur with the following conditions:
- Unable or unwilling to drink water (e.g., illness, injury or experimental or surgical procedures that decrease water consumption for prolonged periods of time)
- Surgical procedures that expose body cavities to evaporative or blood loss
- Clinical conditions that increase water loss (e.g., diarrhea, fever, diabetes, renal disease)
How to Determine if a Rodent is Dehydrated
Skin turgor is used clinically to evaluate dehydration by lifting and releasing the skin of the back over and behind the shoulders. In a well-hydrated animal, the skin almost immediately returns to normal position when released. Skin tenting (delayed return to normal position) can be correlated with degree of hydration. Be aware that skin turgor appears normal in mildly dehydrated (<5%) animals.
If the speed of return to normal position is delayed but the skin does not remain tented, the animal is moderately (~8%) dehydrated. If the skin remains tented for more than 2-3 seconds, the animal is likely severely dehydrated (~10%) and in danger of dying. Obesity and old age may complicate interpretation of skin tenting. Use the overall appearance and clinical history of the animal to aid in making decisions about hydration levels. An ARP veterinarian must be consulted if illness or clinical signs are beyond those expected for the experimental study.
Dehydrated rodents lose weight due to water loss and decreased appetite. Body weight loss over a short period of time (e.g., <48 hours) can be a sign of dehydration and should be treated as such. Other clinical signs of severe dehydration include sunken and dry eyes, listlessness and inactivity, and no urine or fecal output (dry bedding, no feces in cage).
Treatment of dehydration requires administration of fluids orally and/or parentally to correct the animal's fluid deficit and maintain normal hydration after correction. The document below outlines treatment options for dehydration in rodents. Please contact an ARP veterinarian for assistance.
Dehydration and Fluid Therapy in Rodents
Oral Gavage
Liquid compounds may be administered directly into the stomach of rodents via a technique called oral gavage. In this procedure a stainless steel bulb tipped gavage needle or a flexible cannula or tube is attached to a syringe and used to deliver the compound into the stomach. Gavage needles come in various sizes and lengths. The correct needle or cannula length is equal to the distance from the mouth to the last rib. The recommended maximum volume for administration is 1% of body weight (e.g., a 20 gm mouse can be given 0.2 ml).
Correct restraint technique is critical for successful oral gavage. Prior gentle handling to accustom the animal to human hands and restraint is helpful. The animal should be gently restrained (grasp the animal by the loose skin of the neck and back) to immobilize the head but not such that the animal vocalizes or shows other signs of distress. Maintain the animal in an upright (vertical) position and pass the gavage needle along the side of the mouth. Following the roof of the mouth, advance the needle into the esophagus and toward the stomach. If resistance is encountered you may be attempting to enter the trachea and you should alter your needle position. After the needle is passed to the correct length, the compound may be injected. If the animal coughs, chokes or begins to struggle after compound administration begins you may be injecting material into the lungs. If this occurs stop and withdraw the needle immediately. If it appears that material has been injected into the lungs the animal should be euthanized. Struggling during administration or excessive force in advancing the needle may lead to rupture of the esophagus or stomach. If you suspect this has occurred the animal should be euthanized. Sedation or anesthesia prior to oral gavage is not recommended as this will increase the risk of aspiration pneumonia. A video demonstration of oral gavage in the mouse and rat may be viewed on the RAT Procedures With Care website. Oral gavage training videos for mice and rats (produced by Instechlabs) are also available.
Potential Complications
- Passive reflux if excessive material administered.
- Aspiration pneumonia if material injected into the trachea.
- Pharyngeal, esophageal and gastric irritation or injury due to incorrect technique or caustic substances. This may lead to scarring and narrowing of the esophagus and/or gastric openings and esophageal or gastric rupture.
- Physical and/or psychological stress to the animals. Training and habituation to handling may decrease this.
- Microaspiration of material into the lungs (Craig MA, Elliott JF. 1999. Contemp Top Lab Anim Sci 38: 18-23.).
The risk of complications may be decreased by using soft (flexible) gavage tubing rather than stainless steel dosing needles.
References:
Al Shoyaib A, Archie SR, Karamyan VT. Intraperitoneal Route of Drug Administration: Should it be used in experimental animal studies? 2020 Pharm Res 37: 12.
Diehl KH, Hull R, Morton D, Pfister R, Rabemampianina Y, Smith D, Vidal JM, van de Vorstenbosch C. A Good Practice Guide to the Administration of Substances and Removal of Blood, Including Routes and Volumes. 2001. Journal of Applied Toxicology 21, 15-23.
Field KJ, Sibold AL. The Laboratory Hamster & Gerbil. 1999. Boca Raton, FL: CRC Press.
Flecknell PA. Laboratory Animal Anesthesia, 4th ed. 2016. Oxford, UK: Elsevier.
Fox JG, Barthold SW, Davisson MT, Newcomer CE, Quimby FW, Smith AL. The Mouse in Biomedical Research (2nd ed.). 2007. New York, NY: Elsevier Academic Press.
Hankenson FC. Critical Care Management for Laboratory Mice and Rats. 2014. Boca Raton, FL: CRC Press.
Hawk CT, Leary SL, Morris TH. Formulary for Laboratory Animals (3rd ed.). 2005. Ames, IA: Blackwell Publishing.
Hedrich H, Bullock G (Eds.) The Laboratory Mouse. 2004. London: Elsevier Academic Press.
IQ 3Rs Leadership Group - Contract Research Organization Working Group. Recommended Dose Volumes for Common Laboratory Animals. 2016.
Suckow MA, Weisbroth SH, Franklin CL (Eds.). The Laboratory Rat (2nd ed.). 2006. New York, NY: Elsevier Academic Press.
Turner PV, Brabb T, Pekow C, Vasbinder MA. Administration of Substances to Laboratory Animals: Routes of Administration and Factors to Consider. 2011. JAALAS 50 (5): 600-613.