Welcome to the course Writing an Animal Protocol for Research on Mice.

This is the mouse module in a course series on the preparation of an animal use protocol. Each course in this series refers to a different animal species. Every course offers information that is both pertinent to all research animals and specific to the one animal species presented.

• The first 7 lessons are similar in all courses of this series. Differences in these lessons relate mainly to the regulatory coverage, housing requirements, and the zoonotic hazards of each animal species.

• The remaining lessons present information that is more specific to each animal species, such as biological features, anesthetic doses, and biomethodologies.

The goal of this course is to cover important information about using mice in biomedical research settings. If you are responsible for handling mice or if you must write an animal use protocol, this course will be useful by providing you with:

• Information on key regulatory issues.
• Guidance on searches for alternatives in the care and use of animals.
• Highlights of unique biological features of these animals.
• Overviews of acceptable basic methodologies.
• Requirements for supportive care procedures.

Hypertext links in this course provide you with supporting information, such as regulatory sources, drug doses, practical tips, etc.

This course will not provide you with detailed information on how to conduct the methods and procedures described. For this, you should use other courses offering in-depth information and hands-on instruction from your institution's animal facility staff.

View the credits for this course.

To ensure the humane treatment of laboratory animals, animal research is regulated by two federal agencies:

• The United States Department of Agriculture (USDA) / Animal Care; and
• The Public Health Service / Office of Laboratory Animal Welfare.

The USDA and PHS mandates on animal welfare differ greatly with respect to the laboratory strains of mice and rats. These species are not covered by the USDA but are included in PHS regulations and policy. However, the USDA may eventually regulate these species as well.

If your institution receives any funding from the PHS or is accreditated by the Association for Assessment and Accreditation of Laboratory Animal Care, International (AAALAC), then your research must also comply with the National Research Council publication, the Guide for the Care and Use of Laboratory Animals. This document will simply be referred to as the Guide in this course.

The Public Health Service Policy requires institutions to have an occupational health and safety program for individuals working with laboratory animals. This requirement is also reiterated in the Guide.

It is the responsibility of principal investigators to assure that their laboratory staff are informed of and participate in their institution's occupational health and safety program.

Elements of an occupational health and safety program, including institutional responsibilities, are described in the National Research Council publication Occupational Health and Safety in the Care and Use of Research Animals (shown at right).

Working with mice is associated with several types of hazards, which are discussed here and on the next page:

Injuries
Personnel handling a mouse can be bitten if the animal is poorly restrained. Though mice are often inclined to bite when frightened, fortunately their incisors do not always penetrate disposable gloves to break the skin. Bites can be caused by poor handling and restraint technique, which can also cause injury to the mice. If you are nervous working with mice or do not know how to properly handle and restrain them, ask for help. Each instituion must provide training as needed so that personnel know how to handle and restrain mice effectively and humanely, preventing injuries to both people and mice.

Allergies
People can develop an allergy to mice over time after having contact with them. Mouse urine is particularly allergenic, and pelt proteins can also be allergenic. For this reason, you should consider always wearing disposable gloves and a protective gown or scrubs to prevent skin contamination, and a mask to prevent aerosol exposure to urine and pelt proteins. People who develop allergy symptoms should seek medical counseling, and may have to wear special protective equipment or even discontinue working with this species if symptoms are severe after exposure. Protect yourself ! To see some references on allergies to mouse urine, click here.

Zoonoses (diseases transmitted by animals to humans)

In general, transmission of zoonotic disease from naturally infected laboratory animals is uncommon. This is because of ongoing vendor efforts to improve the health status of animals, as well routine periodic infection surveillance programs by facility staff. However, experimentally infected animals are a potential source of zoonotic transmission to humans, and contact with wild mice in field research may also expose humans to zoonotic agents. Animal infection surveillance programs, routine sanitation, training, and personal protective equipment all have important roles in preventing zoonoses.

Mice can be a reservoir of the following infectious agents that are transmissable to people. Here are some zoonotic agents carried by mice:

Viruses

• Hantavirus
  
  

  Hantavirus is a bunyavirus carried by wild mouse species. The virus is transmitted to man by excretions and aerosols from the lungs, saliva, and urine of infected animals.

  Humans are at risk for Hantavirus infection (Korean Hemorrhagic Fever) primarily from wild caught rodents (e.g., the deer mouse, Peromyscus). Strains vary in symptoms based on geographical origin (US, Asia, Scandinavian, and Europe). Hantavirus occurring in the southwestern U.S. causes a severe pulmonary syndrome. Strains orginating in Asia produce a hemorrhagic fever and nephropathy. Strains originating in northern Europe produce renal symptoms of less severity.

• Lymphocytic choriomeningitis virus (LCM)
  
  

  The LCM virus is an RNA arenavirus. Human infection with LCM has been associated with laboratory animals and pets. Mice may be endemically infected (infected in the absence of clinical signs). In utero or early neonatal infection produces a subclinical infection in mice that is characterized by virus shedding (blood, urine). Tumor cell lines may be infected.

  Virus transmission occurs by direct contact as well as by inhalation. Pregnant women are at risk of transmission to the fetus.

  Humans typically develop an influenza-like illness. Additionally, infection may cause a maculopapular rash, lymphadenopathy, meningoencephalitis, orchitis, arthritis, and epicarditis.

Bacteria

• Leptospira spp.
  
  

  Mice may be a reservoir for Leptospira spp. bacteria, which are shed in the urine. Transmission occurs by contact with urine and tissues, or inhalation or ingestion of aerosol droplets.

  Humans with leptospirosis may have influenza-like symptoms, orchitis, rash, skin and mucosal hemorrhage, hemolytic anemia, hepatorenal failure, jaundice, encephalitis, and pneumonia.

• Salmonella spp.
  
  

  Mice may carry Salmonella spp., which are ubiquitous in nature. These bacteria are transmitted via the fecal-oral route.

  Humans infected with Salmonella may have inapparent clinical signs (and be carriers) or may have a febrile enterocolitis, septicemia and focal infections in diverse tissues. Increased severity of the disease occurs due to reduced immunocompetence, e.g., in persons with AIDS, neoplasia, immunosupression therapy, etc., and due to treatment of antibiotics.

Fungi

• Microsporum spp., Trichophyton spp.
  
  

  Dermatophytic fungi grow in the skin and hair follicles and cause a condition of reddened skin and patchy hair loss known as ringworm. The symptoms are the same in animals and humans. Infection may be inapparent in individual animals.

  Dermatophytes are spread by direct contact. Fungal spores are long-lived and may become widely dispersed in the environment. Infections are treatable, but an extended period of therapy is often required to eliminate infection.

Parasites

• Hymenolepsis nana
  
  

  Hymenolepsis nana, otherwise known as the dwarf tapeworm, may be found in mice. It has both a direct and indirect (via flour beetles or fleas) cycle.

  H. nana is transmissable to man. Depending on the parasite burden, humans may have no apparent clinical signs or may have nausea, anorexia, vomiting, diarrhea, and central nervous signs (agitation).

For more information, refer to Occupational Health and Safety in the Care and Use of Research Animals, published by the National Research Council.

Your protocol form should ask you for an assurance that you have considered alternatives to the use of animals if painful or distressing procedures are proposed. This is to satisfy mandates by the Animal Welfare Act and PHS Policy to avoid or minimize discomfort, pain, and distress consistent with sound scientific practices. Alternative procedures are those which may replace animals with nonanimal methods, reduce the number of animals used, or refine the methodology to minimize animal pain or distress. For more information on what is meant by alternatives to the use of animals, please refer to the course Working with the IACUC, which is part of this series.

The assurance often takes the form of a written narrative that describes which sources were used to determine that alternatives were not available. Typically, you may be asked to provide the results of a database search including information on:

1. The databases searched.
2. The date the search was performed.
3. The years of citations covered by database searches.
4. The key words and/or search strategy used when searching a database.

It is strongly recommended that this information be sought during development of a protocol.

Organizations that can assist you in performing an alternatives search are:

• ALTWEB, Center for Alternatives to Animal Testing, Johns Hopkins University
• Animal Welfare Information Center, National Agricultural Library
• Center for Animal Alternatives, University of California - Davis

The following is a simple illustration of how database searches for alternatives can prove very beneficial. The example is intended to help you in the development of a search strategy that is pertinent to your own research.

Click on each database below for a sample search on key terms for this example. You can follow the links to the database for a search in real time.

Example Search:
Determine the alternative models for dietary cholesterol and human cholesterol 7alpha-hydroxylase gene (CYP7A1) in transgenic mice.

PubMed

PubMed Clinical Queries

AGRICOLA

Websites: CYP7A1

More Example Searches:
Additional search routines are available elsewhere in this course to provide you with guidance on performing searches on alternatives in the care and use of animals related to the lesson topic.

The core intent of all of the federal laws, regulations, policies and guidelines applicable to animal research is to ensure the humane treatment of the animals involved in a study. Accordingly, your IACUC will have requirements for the proper care of your animals prior to, during and after a research procedure.

What is a procedure? A procedure is any activity performed on the animal, such as controlled behavioral observation (e.g., use of a maze), venipuncture, or surgery. Requirements for peri-procedural care include:

• Properly preparing the animal to undergo the procedure humanely;
• Supporting the animal's physiological functions during the procedure; and
• Providing appropriate supportive care to aid the animal in recovering from the procedure.

The investigator has the responsibility to see that staff working with the animals are properly trained not only to perform the procedure humanely but also to provide the necessary supportive care to the animals.

When performing any procedure, you should think through the steps that are necessary to protect the animal's welfare. For example, for blood collection, you should limit the volume taken to a safe minimum and you should realize that safe volumes will differ for acute or chronic collections. With any venipuncture, you should be prepared to care for the animal in the event of trauma to the vein or excess hemorrhage.

The saphenous vein shown in the photo is useful only for small volume collections.

Refer to your institution's IACUC and attending veterinarian for specific guidelines.

Your protocol form may ask you which type of housing you may need for your mice. There are important considerations in the selection of animal housing that affect the welfare of your animals.

Rodent caging has two types of flooring: solid and wire mesh.

Because of data on rodent preferences for solid flooring and the risks for animal injury on wire mesh flooring, the use of wire bottom cages should be scientifically justified and approved by your institution's IACUC. Related guidelines are:

Guide for the Care and Use of Laboratory Animals

ARENA/OLAW IACUC Guidebook

Example Alternatives Search:
For additional information on types of mouse caging and the impact of caging type on mice, please refer to the examples of "alternatives" searches on mouse housing.

Upon arrival to your facility, your mice should have an acclimation period before they are used in research studies. This period of time allows animals to adapt to a new environment. Effects of transportation stress include alterations in various blood parameters, immune cell function and animal behavior. The period of time necessary for biological stabilization will depend on the parameters to be studied. Refer to your institution's attending veterinarian for recommendations that are appropriate for your project. Typically, acclimation periods can range from days to over a week, depending on the studies involved.

Example Alternatives Search:
For citations on the impact of transportation stress and acclimation on mice, please refer to the examples of "alternatives" searches provided.

Routine quarantine procedures may prolong the holding of your animals in special facilities. An important goal of quarantining animals is to prevent transmission of diseases between new animals and animals already present at the facility in established colonies.

Many institutions quarantine all mice received from other institutions, no matter what certifications of health may accompany them. There are many reasons for this, but the following three are worth noting:

1. Detecting viral, bacterial, and parasitic pathogens in mice can be challenging because many infections are asymptomatic (cause no observable clinical signs), and thus infections can be missed in animals prior to shipment. As an example, pinworm infections in mice are notoriously difficult to diagnose because eggs from the female nematodes are shed intermittently and sometimes in low number, leading to missed diagnoses.

2. The cost of controlling and eliminating infections once they escape into other colonies can be enormous.

3. And finally, huge amounts of investigator time as well as priceless research data can be lost due to infections.

Acclimation and quarantine periods can run concurrently, although they serve different purposes. Institutions may or may not allow experiments on animals while quarantined, depending on the circumstances.

If your proposed study involves a painful procedure, the protocol form may ask for a method of assessing if the mice are experiencing pain or distress.

Assessing pain and distress in mice is difficult at times because mice, like many other species, commonly conceal outward signs of moderate pain and distress. Accordingly, behavioral changes that reveal a mouse's pain and distress may be subtle and elude detection unless observations are thorough and made by a trained observer.

Severe pain and distress causes overt clinical signs in mice. Laboratory staff working with mice should be trained to recognize these abnormalities in:

• Activity level: hypoactivity (abnormally low), hyperactivity (abnormally high), restlessness.
• Behavior: vocalization, self-trauma, aggressiveness, isolation from cage mates, ataxia.
• Appearance: unkempt or greasy fur, porphyrin staining around eyes and nostrils, hunched posture, cyanosis, pale mucous membranes, soiled anogenital area.
• Vital Signs: e.g. respiratory distress.
• Body Condition: weight loss, emaciation, dehydration.
• Intake: reduced intake of food and water.

The mouse shown above right has scruffy fur, a hunched posture, and porphyrin staining around the orbit. The ears, feet, and tail have a blanched coloration, suggesting vasoconstriction (blood vessel constriction) or hypoperfusion (abnornally low levels of blood in tissue). This mouse is showing signs of severe pain and/or distress.

A chronic state of pain or distress may be more subtle and difficult to detect. A good knowledge of the animal’s normal appearance and behavior is especially important to recognize chronic pain or distress.

For methods on assessing and alleviating pain and distress in rodents, refer to another course in this series, Post Procedure Care of Mice and Rats in Research: Minimizing Pain and Distress.

Example Alternatives Search:
For citations on detecting pain and distress, please refer to the example of "alternatives" searches provided.

Breeding mice as inbred strains and outbred stocks produce animals that are used for different purposes. The decision to use isogenic inbred strains or non-isogenic outbred stocks is determined by the experimental strategy.

Inbred strains are used for genetic engineering and finely controlled studies that capitalize on genetic isogenicity. Inbred strains with characteristics of human diseases or physiological conditions are generally preferred models for biomedical research.

Outbred mice are used when outbred vigor is desirable, e.g., as foster females for a transgenic colony, or when genetic heterogeneity and phenotypic variability are not a concern.

Please check with your animal resource department for information on vendor choices as animal source affects animal health status.

Though mice share many anatomical and physiological features with humans, mice have many unique biological characteristics. A knowledge of species-specific characteristics is helpful to effectively manage these animals and to plan experimental procedures for their use.

Researchers should be aware of the following practical features of mouse anatomy and biology:

Anatomy

• Ocular System
  
  Rats and mice may develop red staining around the eyes and nostrils when they are distressed, e.g., by disease, trauma, etc. This staining is due to the accumulation of porphyrins produced by the Harderian gland, a lacrimal gland. Though a normal constituent of tears in rodents, lacrimal porphyrin is produced in limited amounts and rodents keep themselves clean of debris through frequent grooming. Porphyrin staining in distressed animals occurs because stress stimulates porphyrin production in tears and distressed animals groom themselves less often.



• Teeth
  
  Mice have incisors that are open rooted, meaning that these teeth grow continuously throughout adult life. A diet of soft foods, i.e. in liquid or powder form, or a developmental jaw malformation will cause tooth overgrowth. In particular, transgenic or knock-out mice may have unintended genetic anomalies that cause jaw malalignment and result in tooth overgrowth. Staff must be alert to detect any signs of this condition and to provide appropriate treatment.

Gastrointestinal System

• Inability to vomit
  
  Mice do not vomit because they lack the neurophysiological mechanisms for doing so. Therefore, withholding food and water before surgery is not usually necessary in mice.



• Gall Bladder
  
  Unlike rats, mice do have a gall bladder.



• Coprophagy
  
  In mice, herbaceous foodstuffs are broken down by microbial action in the cecum, which is a large organ in the mouse. To assimilate the microbial byproducts of digestion, the mouse regularly eats its own feces, a habit known as coprophagy. If a study does require fasting for scientific reasons, be aware that mice will consume their own feces and thus there may be fecal material in the GI tract in the absence of food.Stomach digestion and intestinal absorption of this fecal material yields nutrients that are essential to the mouse.

Metabolism

• Albinism
  
  Most mice used in research are albinos, whether an inbred strain such as the Balb/C or an outbred stock such as the Swiss Webster. Albinism in mice is an inherited disorder of melanin metabolism caused by the lack of the enzyme tyrosinase, which has an impact both on melanocytes and neurons. Neuronal morphological abnormalities and functional impairments involve the following sites: medial vestibular nucleus, cochlear nuclei and retina. Studies comparing albino and pigmented animals have shown differences even in pharmacotoxic kinetics in these tissue areas. The lack of pigment in the eyes of albinos can result in retinal damage in brightly lit caging rooms. Consequently, animal care staff are obligated to monitor light levels.



• High rate of metabolism – impact on drug clearance


The mouse’s high rate of metabolism produces a rapid clearance of drugs from the body. Drugs administered at dose rates used in larger species (with lower metabolic rates) will likely reach lower blood concentrations and exert less pharmacological effect in the mouse. This includes analgesics given postoperatively to control pain. As a result, mice should receive drug doses that have been scaled to the mouse’s metabolism. Through a discipline known as allometry, mathematical formulas have been developed to adjust dose rates between species of disparate size.

In general, mouse-specific dose rates have been determined and are widely published for drugs that are commonly used in animal research, such as anesthetics, analgesics, sedatives, and antibiotics. Investigators are advised to obtain mouse dose rates from laboratory animal references or from their institution's veterinary staff.


• High surface area – impact on hypothermia


Mice have a large body surface area (relative to body volume) plus many hairless body parts (tail, ears, feet). As a result, mice are vulnerable to profound hypothermia under conditions of sedation and anesthesia. Sedation and anesthesia induce hypothermia due to drug effects on the hypothalamus and a rapid drop in core body temperature. If surgery is being performed, additional heat is lost by convection from an open incision during surgery, and placement of the mouse on a heated surface may be necessary during the surgery to maintain a healthy body temperature.

Mice should have a source of warmth throughout a procedure that lowers their body temperature (e.g., anesthesia, surgery) and afterwards until they recover the ability to thermoregulate themselves.

Volume recommendations (ml) for acute intravenous fluid administration and blood collection in adult mice (average 20 g):

IV Fluid Volume (ml) max. acute admin.	Total Blood Volume (ml)	Safe Bleeding Volume (ml)a	Tot. Bleed-out Volume (ml)b
0.2	1.0 - 2.4	0.1 - 0.2	0.6 - 1.4

^aRemoving greater quantities of blood (exceeding 0.1 ml per 10 grams of body weight, or alternately expressed, about 10% of total blood volume) can produce hypovolemic shock. Repeated collections of smaller amounts of blood will have the same effect. In such procedures, it may be necessary to administer warmed physiological fluid to replace the volume of blood collected.

^bAnimals should be exsanguinated only under anesthesia.

(From Wolfensohn and Lloyd, Handbook of Laboratory Animal Management and Welfare, 2nd Edn., Blackwell Science. 1998.)

Below are peripheral vessels that are commonly accessed for blood collection or fluid administration. Recommended needle sizes are 25 to 29 gauge. Larger needles may be necessary for injecting large volumes or viscous materials.

Below are the nonvascular routes of injection that are commonly used in mice. Included are volume recommendations for the safe administration of fluids acutely in adults (average 20 g). Recommended needle sizes are 25 to 27 gauge; larger needles may be necessary for injecting viscous materials.

Subcutaneous (SQ or SC)
2-3 ml total; maximum of 0.5 ml per site.

Intraperitoneal (IP)
2-3 ml

Oral (PO)
0.4 ml

Intradermal (ID)
0.05 ml/site

Note –
Intramuscular (IM) injection is not generally recommended in mice because these animals lack sufficient muscle mass for an injection. An IM injection in mice would be likely to cause muscle injury. If an IM injection were necessary, the volume administered should not exceed 0.05 ml per site.

Example Alternatives Search:
For citations of blood collection procedures in mice, please refer to the examples of "alternatives" searches provided.

The in vivo method of monoclonal antibody production uses mice to grow hybridoma cells on the peritoneal lining of histocompatible animals. Monoclonal antibodies are then collected from the antibody-rich ascites fluid.

Over the past years, there have been a number of in vitro techniques introduced that can replace the use of animals for expanding hybridoma cell lines and collecting purified monoclonal antibody. Consequently, non-animal alternatives for generating purified monoclonal antibodies should be considered. The in vitro method should be considered and deemed unsuitable on scientific grounds before the IACUC approves animal use for the in vivo method.

When requesting approval to use animals for expanding hybridoma cell lines, be prepared to explain why in vitro techniques will not work. In 1999, The Committee on Methods of Producing Monoclonal Antibodies (sponsored by the Institute for Laboratory Animal Research and the National Research Council) suggested the following guidelines for IACUCs to use when evaluating the need for using animals for hybridoma expansion (Recommendation 4):

-When a supernatant of a dense hybridoma culture grown for 7–10 days (stationary batch method) yields a monoclonal antibody concentration of less than 5 mg/ml, or if other systems are used and concentrations obtained are less than 500 mg/ml (hollow fiber system) and 300 mg/ml (semi-permeable membrane system).

-When more than 5 mg of monoclonal antibody produced by each of five or more different hybridoma cell lines is needed simultaneously. It is technically difficult to produce this amount of monoclonal antibody since it requires more monitoring and processing capability than the average laboratory can achieve.

-When analysis of monoclonal antibodies produced in tissue culture reveals that a desired antibody function is diminished or lost.

-When a hybridoma cell line grows and is productive only in the animal.

-When more than 50 mg of functional monoclonal antibody is needed, and previous poor performance of the cell line indicates that hollow-fiber reactors, small-volume membrane-based fermentors, or other techniques cannot meet this need during optimal growth and production.

These same criteria can help you decide if in vitro methods will suffice. The burden of proof is now on the investigator to show that in vitro methods of obtaining purified monoclonal antibody do not work, or are not effective in providing the amount of antibody needed.

Example Alternatives Search:
For consideration of the alternatives to using mice for the production of monoclonal antibodies, please refer to the examples of "alternatives" searches provided.

If in vivo methods are needed because in vitro methods cannot replace them, consideration must be given to minimizing the amount of pain and suffering involved. The following parameters should be considered when animals are used to expand hybridomas using the ascites collection technique:

1. The amount of pristane used to “prime” the peritoneal cavity and make it better able to support hybridoma growth should be minimized.
2. The degree of abdominal distension should be monitored at least daily and should distension begin to interfere with breathing, the ascites fluid should be removed.
3. The number of peritoneal “taps” used to collect ascites fluid should be minimized.
4. The needle used should be as small as possible (20 gauge or higher). Because mice with ascites are not good anesthetic risks, ascites fluid is usually collected with a needle and syringe without anesthesia, and smaller bore needles cause less pain.
5. Endpoint criteria tailored to collecting ascites should be developed. In addition to typical endpoint criteria such as weight loss and extended anorexia, additional criteria to consider include a limit on the number of abdominal taps allowed, the presence of dyspnea (difficult breathing) unrelieved by a tap, and the development of solid hybridomas instead of more diffuse neoplasms producing ascites. Usually you will be asked about endpoint criteria somewhere on your forms.

If you anticipate the need for adjuvants to stimulate an immnue response, several issues must be considered.

The classic immune adjuvant is Freund's adjuvant, which is available in two forms:

• "Complete" (Complete Freund's Adjuvant, or "CFA"). CFA is a mixture of oils and water plus killed Mycobacterium tuberculosis. It typically elicits a very strong immune reaction. If used more than once, the immune reaction usually progresses to intense inflammation and sterile abscesses.
• "Incomplete" (Incomplete Freund's Adjuvant, or "IFA"). IFA is similar to CFA, but is missing the killed mycobacteria. This renders the IFA less effective as an immune stimulant, but also less inflammatory to the animal. IFA can be used safely multiple times without causing intense inflammation.

To prevent inflammation and pain, CFA must be used once only. IFA is less inflammatory, and can be used multiple times. Typically CFA mixed with antigen is given to an animal the first time, then IFA mixed with antigen is given second, then either IFA mixed with antigen or antigen alone for subsequent immunizations.

The USDA states that the injection of CFA may cause more than momentary or slight pain. This means that CFA injections might put an animal into USDA pain category D (painful/stressful but relieved), requiring the use of post-injection analgesics or sedatives.

You may be asked to list any possible medical complications resulting from the use of adjuvants during polyclonal antibody production in your animal protocol forms.

To reduce inflammation when using CFA, consider the following measures:

1. Choose or make preparations of CFA with a lower mycobacterial concentration, i.e., 0.05 to 0.1 mg/ml, rather than 1 mg/ml.
2. Add a concentrated antigen solution to the adjuvant to obtain a more antigen-rich emulsion, thereby reducing the volume of emulsion injected.
3. Use multiple injection sites to limit the volume injected at any one site.
4. Separate the injection sites to avoid fusion of inflammatory lesions.
5. Maintain sterility of the antigen solution.

The quantity of CFA or IFA adjuvant injected should be limited. Institutional guidelines vary, but typical limits on adjuvant use are around 0.25 to 0.50 ml combined adjuvant/antigen per immunization for smaller animals such as mice (up to ten divided 0.05 ml injections). These amounts have been shown to produce high titer antibodies, yet limit inflammation. Check with your institution for specific guidelines in place.

Beware: CFA is a health hazard to humans.

If you are already sensitized to mycobacterial antigens by a previous exposure to CFA, immunization against tuberculosis, or through a natural infection of tuberculosis, you are likely to experience severe inflammation if you splash CFA into your eye or accidentally inject yourself with it. The inflammation and pain may be so severe that surgical removal of the site may be necessary. Protect your eyes and prevent accidental injection of yourself or a colleague when using CFA!

Less inflammatory alternatives to CFA and IFA are now available and in use. Examples are the block copolymer adjuvant Titermax®, and the lipid A-derivative adjuvant MPL® by RIBI. Other promising alternative adjuvants are also on the market. Such alternatives can be considered as a means of further reducing inflammation induced by Freund`s adjuvant.

The route of immunization should be chosen to limit pain and inflammation. Regardless of the adjuvant used, the subcutaneous route typically provides a strong immune response, and is recommended. The intravenous route is not appropriate if adjuvant is used because the thick consistency of the adjuvant can result in lethal emboli in the blood stream.

There are several other routes of adjuvant immunization that are usually discouraged, unless there is clear evidence that they offer an advantage over the subcutaneous route for a specific use:

Intradermal (ID): Causes more pain because the skin itself cannot stretch much as body fluids and white blood cells enter the immunization area, resulting in increased pressure and pain. Some institutions allow the ID route if less volume of adjuvant and antigen are injected.

Intraperitoneal (IP): Inflammation on surfaces of abdominal organs can result in peritonitis, granulomas, and pain.

Foot pad: Injections can cause pain and lameness. When allowed by an IACUC, usually only one foot may be injected. Foot pad injections are usually discouraged in rodent species, and deemed inappropriate in larger species.

Because mice have a high rate of metabolism, drugs are rapidly eliminated from their bodies. Dose rates appropriate for larger species produce ineffective drug concentrations when used in mice.

This section includes mouse dose rates for the common drugs and drug regimens. If you need to use other drug agents, check with your institution's veterinary staff for assistance in determining a dose rate appropriate for use in mice.

Click on the drug types for doses of common agents and drug regimens that may be used in mice:

Analgesics:
Available in two drug types – the opioids and the nonsteroidal anti-inflammatory drugs (NSAIDs). The rapid clearance of many of these drugs in mice results in the need for an increased frequency of administration.

Sedatives:
Sedatives may obtund consciousness but in normal doses do not do so sufficiently to ablate the perception of pain or other sensations. When combined with general anesthetics, they may be used to induce a "balanced" anesthesia where muscle relaxation, unconsciousness, and analgesia are enhanced.

Sedatives + Analgesia:
Some sedatives also have analgesic effects. When combined with general anesthetics, these sedatives enhance analgesia and a "balanced" anesthesia is attained.

Anesthetics:
Because mice metabolize drugs so rapidly, many anesthetic agents have brief durations of effect. An anesthetic regimen should be chosen to match the duration of drug effects with the length of the procedure. In particular, short acting agents (and regimens) should be not be used for long procedures because repeat drug administrations, necessary to prolong anesthesia, will produce uneven blood concentrations and therefore periodically inadequate anesthesia. For long procedures, gaseous anesthesia using a non-explosive agent such as isoflurane is often the most practical method to sustain uniformly adequate levels of anesthesia. Potentially explosive agents, such as ether, are not recommended.

The practice of using hypothermia as an anesthetic for neonates is generally discouraged.

It is not clear whether the depression of neural function by hypothermia is sufficient to prevent the sensation of pain related to a surgical procedure. Also, the recovery from hypothermia may be a painful experience in animals, as it is known to be in humans.

Inhalation anesthesia with an agent such as isoflurane administered using a non-rebreathing system may be an acceptable alternative to hypothermia in neonatal rodents.

Example Alternatives Search:
For additional information on anesthesia in mice, including the physiological impact of anesthetic agents on mice, please refer to the examples of "alternatives" searches provided.

Aseptic technique should be used when performing surgery on mice. The standards described here are consistent with the Guide for the Care and Use of Laboratory Animals.

Surgery on mice should be performed in a location that allows for a physical separation of the operative field from other functions of the procedure (such as animal preparation and anesthetic recovery) and other laboratory activities.

• The isolation of the operative field avoids contaminating sterile areas with animal fur, bedding, nonsterile supplies, etc.
• The location used for the operative field should be cleaned and sanitized before use.
• Materials and supplies used in support of the procedure should be positioned and managed to avoid contaminating sterile areas.

Surgical procedures in mice should be conducted using aseptic technique. Nonaseptic methods are not acceptable. Rodents have been shown to develop subclinical infections, a consequence which has led to an outdated belief that rodents tolerate nonaseptic technique without developing postoperative infections. The Guide recommends methods for adapting aseptic technique to the scale of rodent surgery. In this way, efficiencies and economies can be realized without sacrificing asepsis.

Supportive care aims to:

• Maintain the animal's physiological status as nearly normal as possible.
• Minimize animal pain and distress.

Supportive care includes monitoring of both physiological parameters and analgesia during anesthetic and surgical procedures. Monitoring of vital signs and pain perception should be conducted throughout the procedure and the recovery period.

Keep in mind that:

• General anesthesia causes disturbances of thermoregulation and other physiological functions. Maintaining body temperature, e.g., via insulating materials and supplemental heating sources, is an important objective of supportive care.
• During surgery, the animal may experience pain if anesthesia is inadequate at any time during the procedure.
• Postoperatively, the animal may experience pain, discomfort, and distress unless treated with analgesics and appropriate supportive care measures.

Due to the interaction of metabolic factors and drug effects that can cause animal mortality, mice should receive good supportive care and monitoring during anesthesia, whether or not the procedure involves surgery.

During anesthesia and surgery, the following procedures are recommended.

Supportive Care:

• Provide a source of warmth to mice from the onset of anesthesia to the end of anesthetic recovery. Care needs to be taken to avoid heating sources that may cause thermal injuries to the mice.
• Inject sterile physiological fluid (warmed to body temperature) to compensate for blood loss during a procedure and depressed fluid intake post-procedure.

Monitoring during Anesthesia:

• Analgesia - toe pinch.
• Respiration -gross changes in rate, character of breathing.
• Color of mucous membrane and skin – blue (poor oxygenation), pale (poor blood perfusion).

After anesthesia and surgery, the following procedures are recommended for:

Monitoring post Anesthesia:

• Mice must be monitored until fully recovered from anesthesia as indicated by the ability to ambulate and maintain core body temperature. Routine use of antibiotics is not indicated after uncomplicated, aseptic surgery.

Monitoring Post Procedure:

• Assess appearance, activity, and behavior as indications of pain and discomfort (see screen Detecting Pain and Distress).
• Assess food and water intake.
• Provide floor-level access of food and water post procedure if stretching overhead for these items (in the cage wirelid) may be painful.
• Assess wound repair.

The term euthanasia is derived from Greek and means "good death." Animals should be euthanatized when killed for any purpose, including research. To euthanatize a mouse, you must be trained in the concepts of euthanasia, the method to be used, and the proper handling of mice.

Methods are classified as acceptable or conditionally acceptable, as set by the American Veterinary Medical Association. The inclusion of conditionally acceptable methods in your protocol may require scientific justification and IACUC approval.

The photo shows a CO₂ chamber for euthanatizing rodents.

Click on each method below for recommendations on its use in euthanatizing mice.

Acceptable Methods:

• Barbiturates
• Inhalant anesthetics
• Carbon dioxide (compressed tanks only)
• Carbon monoxide
• Potassium chloride in conjunction with general anesthesia
• Microwave irradiation

Conditionally Acceptable Methods*:

• Ether
• Nitrogen
• Argon
• Cervical dislocation
• Decapitation

* The inclusion of conditionally acceptable methods in your protocol may require scientific justification and IACUC approval.

Very Important!

Before placing euthantized rodents in a bag and placing the bag in a necropsy refrigerator or freezer, you must make sure the mice are really dead! Mice can stop breathing for a minute or more then regain respiratory function and survive. This is particularly true of younger mice, which are somewhat resistant to carbon dioxide asphyxiation and take longer to succumb than adult mice.

To ensure death in mice euthanatized with carbon dioxide, the chest cavity may be opened with scissors, or the mice may be observed for an extended period of time to make sure they are dead. Your institution may have specific requirements.

The Office of Laboratory Animal Welfare (responsible for enforcing PHS policy) has made it clear that rodents remaining alive in bags after ineffective euthanasia is a serious breach of PHS policy, and must be reported to regulatory officials.

Example Alternatives Search:
For additional information on euthanasia of mice, including the impact of euthanasia agents on tissues, please refer to the examples of "alternatives" searches provided.

Federal Laws, Regulations, Policies:

1. Animal Welfare Act, as Amended (7 USC, 2131-2156)
2. Animal Welfare Act Regulations and Standards, Code of Federal Regulations, Title 9, Part 2-Regulations, Sections 2.31-2.33, 1998.
3. Health Research Extension Act of 1985, Public Law 99-158, November 20, 1985, "Animals in Research".
4. Public Health Service Policy on Humane Care and Use of Laboratory Animals, Revised September, 1986, Reprinted March, 1996.
5. USDA Animal and Plant Health Inspection Animal Care Policy Manual.
   Policy #11 – Painful/Distressful Procedures.
6. U.S. Government Principles For The Utilization And Care Of Vertebrate Animals Used In Testing, Research, And Training, Interagency Research Animal Committee.

Guidelines:

1. Guide for the Care and Use of Laboratory Animals, National Research Council, 1996.

Texts:

1. Hawk and Leary, Formulary for Laboratory Animals. 2nd Edn., Iowa State University Press, 1999.
2. Hrapkiewicz, Medina, and Holmes, Clinical Laboratory Animal Medicine: An Introduction, 2nd Edn., Iowa State University Press, 1998.
3. Suckow, Danneman, and Brayton. The Laboratory Mouse. CRC Press, 2001.
4. Wolfensohn and Lloyd, Handbook of Laboratory Animal Management and Welfare. 2nd Edn., Blackwell Science. 1998.