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

This is the hamster 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 hamsters in biomedical research settings. If you are responsible for handling hamsters 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.

Click to view the credits for this course.

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

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

If your institution receives any funding from the PHS or is voluntarily 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: National Research Council. 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 guideline, Occupational Health and Safety in the Care and Use of Research Animals (shown at right), published by the National Research Council.

Working with hamsters is associated with the following hazards:

Injuries
Personnel handling hamsters can be injured by bites from the incisor teeth. Bites often occur because of a lack of knowledge in how to handle, transport and restrain a hamster. A hamster will bite when startled from sleep, or when frightened, or in pain. Likewise, poor technique in handling, etc., can cause injury to the hamster. Training staff to work effectively and humanely with hamsters is essential to prevent injuries to people and hamsters.

Allergies
People can develop an allergy to hamsters after having contact with them for some time. Persons who develop allergic symptoms should seek medical counseling and may have to discontinue working with this species.

Zoonoses

In general, transmission of zoonotic disease from laboratory animals is uncommon because of the ongoing efforts to improve the health status of animals by vendors of purpose bred animals and by facility staff. Experimentally infected animals are a source of zoonotic transmission to humans. Health surveillance programs, routine sanitation, and personal protective equipment have important roles in preventing zoonoses.

Hamsters can be a reservoir of the following infectious agents which are transmissable to people:

Viruses

Lymphocytic choriomeningitis virus (LCM)

 

Bacteria

Leptospira spp.

 

Protozoa

Giardia spp.

 

Helminths

Hymenolepsis nana

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 must take 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 case study of alternatives searches that may guide 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:
Study of the effect of photoinhibition on the morphology and function of pituitary lactotropes in hamsters.

PubMed

PubMed Clinical Queries

Agricola

BIOSIS

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.

All of the federal laws, regulations, policies and guidelines applicable to animal research have as their core intent 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 carried out on the animal, such as venipuncture or surgery. This peri-procedural care requirement includes:

• Properly preparing the animal to humanely undergo the procedure;
• Supporting the animal's physiological function during the procedure; and
• Providing additional 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 to not only perform the procedure humanely but also to provide the necessary supportive care to the animals.

When performing any procedure, such as a blood collection, 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 to the minimum that may be taken safely. That amount may vary depending upon whether a single sample is taken or multiple, chronic collections are done and, with any venipuncture, you should be prepared to care for the animal in the event of trauma to the vein area or excess hemorrhage.

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 hamsters. There are important considerations in the selection of animal housing that affect the welfare of your animals. For the appropriate cage space per animal, refer to the minimum requirements specified by the Animal Welfare Regulations.

Hamster caging has two types of flooring: solid and wire mesh. Hamsters may be housed in microisolator cages (filter cage top and solid bottoms) under barrier conditions or in open-topped cages (no filter top) under conventional conditions. Hamster caging must be secure, e.g., with tightly fitting wire lids, because hamsters are prone to dislodge cage tops and escape through narrow spaces. Hamsters have a solitary nature and so may prefer to be housed individually.

The solid flooring of shoebox cages are covered with a bedding material that absorbs liquid wastes. Bedding has been shown to be preferred by rodents for resting, and it is considered to provide them with comfort, warmth, and the opportunity to burrow. This type of flooring is well suited to breeding because pups are better protected from chilling.

 

Wire mesh flooring offers advantages in sanitation. Wire spacing must be narrow enough so that the hamsters' feet do not slip through the holes, which could injure the feet. The use of wire bottom cages is discouraged for rodents, especially on long-term studies.

Because of data on rodent preferences for flooring type, the Guide for the Care and Use of Laboratory Animals recommends the use of cages with solid flooring for rodents. The use of wire bottom cages should be scientifically justified and approved by the IACUC.

Example Alternatives Search:

For additional information on hamster housing, please refer to the example "alternatives" search on hamster housing.

Upon arrival at your facility, your hamsters 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 range from 4 days to 1 week.

Example Alternatives Search:
For hamster related citations on acclimatization, including transportation stress, please refer to the example of "alternatives" searches provided.

Routine quarantine procedures may prolong the holding of your animals in special facilities. Quarantine aims to prevent transmission of diseases between new animals and established colonies.

Acclimation and quarantine periods run concurrently, although they serve different purposes. Most institutions do not allow experiments on animals while quarantined.

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

Assessing pain and distress in hamsters is difficult at times because hamsters, like many other species, commonly conceal outward signs of moderate pain and distress. In this case, the behavioral changes that reveal a hamster's pain and distress may be subtle and detectable only with specialized behavioral methods.

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

• Activity level – e.g., depression, or extended sleep.
• Behavior – e.g., aggressiveness, self-trauma, ataxia, vocalization (chatter or screech).
• Appearance – e.g., ruffled fur, ocular discharge, hunched posture, cyanosis, pale mucous membranes, soiled anogenital area, diarrhea.
• Vital Signs – e.g., respiratory distress, hyperthermia, hypothermia.
• Body Condition – e.g., weight loss, emaciation, dehydration.
• Intake – e.g., reduced intake of food and water.

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. The methods described are pertinent to all laboratory rodents including hamsters.

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

The hamster species that have been used in research are as follows. The first two are most common.

• Golden hamster or Syrian hamster, Mesocricetus auratus. (adults 120 g)
• Chinese or Striped hamster, Cricetulus griseus (adults 30-35 g)
• Armenian hamster, Cricetulus migratorius (approx. 45 g)
• European hamster, Cricetus cricetus (large)
• Siberian or Djungarian hamster, Phodopus sungorus (adults 30-50 g)

The image below shows a Chinese hamster (left), an Armenian hamster (middle), and a Syrian hamster (right).

Most hamsters are outbred stocks, meaning that individuals have a high degree of heterozygosity and phenotypic variability.

Inbred (isogenic) strains of the Syrian hamster (Mesocricetus auratus) are used for cardiomyopathy studies. These strains are designated:

• CHF148 (spontaneous disease model)
• CHF146 (control)

Inbred strains are typically used for 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.

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

Though hamsters share many anatomical and physiological features with humans, hamsters 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 hamster anatomy and biology.

In the image at right, a hamster's cheek pouch is everted to show the mucosal surface.

Items on the following screens offer a brief description of some unique features and some practical tips.

Behavior

Biting

Escapism

Feeding behavior

Nocturnalism

Social characteristics

 

Anatomy

Cheek pouches

Teeth and caries

Scent glands

 

Metabolism

High rate of metabolism – impact on drug clearance

High surface area – impact on hypothermia

Pseudohibernation

 

Gastrointestinal

Inability to vomit

Coprophagy

 

Immune System

T-cells

Cheek pouches

Volume recommendations for intravenous fluid administration and blood collection in adult Syrian hamsters^a:

IV fluid volume (ml) max. acute admin.	Tot. blood volume (ml)	Safe bleed volume (ml)a	Bleed-out volume (ml)b
0.3 ml	78 ml/kg	0.55 ml	2.9-5.2 ml

^a Removing greater quantities of blood (exceeding 10% of total blood volume) can produce hypovolemic shock. Repeated collections of smaller amounts of blood will have the same effect. In such procedures, animals should receive warmed, physiological fluids to replace the volume of blood collected. In addition, monitor the animal’s hematocrit for anemia.

^bAnimals should be exsanguinated only under anesthesia.

From:

1. Hawk and Leary, Formulary for Laboratory Animals, 2nd Edn., Iowa State University Press, 1999.
2. 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 23 to 27 gauge. Larger needles may be necessary for injecting large volumes or viscous materials.

Vessels	Comments
Lateral saphenous vein (lateral tarsal vein)	Accessing the lateral saphenous vein: Does not require anesthesia. May be aided by sedation because vein visibility is enhanced by peripheral vasodilation (drug effect). May be aided by sedation to reduce animal struggling due to distress. Blood collection from the lateral saphenous vein does not involve cannulation of the vein lumen. Instead, the vein is punctured percutaneously and blood is passively collected as it pools on the skin.
Jugular vein	Performed under anesthesia because of restraint method and the need for animal immobilization.
Cardiac puncture Carotid artery	These two methods require anesthesia. Most often allowed only as a terminal procedure. Check with your institution for guidelines on these routes of blood collection.
Abdominal vena cava Abdominal aorta	Performed as a terminal procedure under anesthesia. Vessel access involves a ventral midline incision and reflection of intestines.
Retroorbital sinus	Retroorbital puncture is controversial because of the risk of injury to the optic nerve and other nearby structures. This method is considered to be painful and may cause blindness. Generally requires anesthesia. Topical ophthalmic anesthetic is recommended post-procedure.

Below are the nonvascular routes of injection that are commonly used in hamsters. Included are volume recommendations for the safe administration of fluids acutely in adults (Syrian hamster, average 120 g). Recommended needle sizes are 23 to 27 gauge; larger needles may be necessary for injecting viscous materials.

Subcutaneous (SQ or SC) - 3-4 ml in scruff

 

Intraperitoneal (IP) - 3-4 ml

 

Oral (PO) - 20 ml/kg

 

Intradermal (ID) - 0.05 ml/site

 

Intramuscular (IM) - 0.1 ml per site

 

Cheek pouch - 0.1 ml instilled into wall of everted pouch.

From:

1. Hawk and Leary, Formulary for Laboratory Animals, 2nd Edn., Iowa State University Press, 1999.
2. Field and Sibold, The Laboratory Hamster & Gerbil, CRC Press, 1999.
3. Wolfensohn and Lloyd, Handbook of Laboratory Animal Management and Welfare, 2nd Edn., Blackwell Science. 1998.

Example Alternatives Search:

For citations of injection and blood collection methods in hamsters, please refer to the examples of "alternatives" searches provided.

Because hamsters 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 hamsters. This section includes dose rates for the common drugs and drug regimens used in hamsters. 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 hamsters.

Click on the drug types for doses of common agents and drug regimens:

Analgesics:
Available in two drug types – the opioids and the nonsteroidal anti-inflammatory drugs (NSAIDs). The rapid clearance of many of these drugs in hamsters 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, a balanced anesthesia is attained, and these sedatives enhance analgesia through specific effects.

Anesthetics:
Because hamsters 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 is often the most practical method to sustain uniformly adequate levels of anesthesia.

For additional information on analgesics, sedatives, and anesthetics in hamsters, including the physiological impact of these agents, please refer to the examples of "alternatives" searches provided.

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

Surgery on hamsters 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 hamsters 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 the monitoring of both physiological parameters and analgesia during anesthetic and surgical procedures. Monitoring of vital signs and pain should be conducted throughout the procedure and the recovery period.

Keep in mind that:

• General anesthesia causes disturbances of thermoregulation and other physiological functions.
• 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, hamsters 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 hamsters from the onset of anesthesia to the end of anesthetic recovery. Care should be taken to avoid use of devices that may cause thermal injury.
• 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 – poor oxygenation (blue), poor blood perfusion (pale)

After anesthesia and surgery, the following procedures are recommended.

Monitoring post Anesthesia:

• Hamsters must be monitored until fully recovered from anesthesia as indicated by the ability to ambulate and maintain core body temperature.

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. Routine use of antibiotics is not indicated after uncomplicated aseptic surgery.

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 hamster, you must be trained in the concepts of euthanasia, the method to be used, and the proper handling of hamsters.

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.

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 animals are really dead! Rodents can stop breathing for a minute or more then regain respiratory function and survive. This is particularly true of younger rodents, which are somewhat resistant to carbon dioxide asphyxiation and take longer to succumb than adult rodents.

To ensure death in rodents euthanatized with carbon dioxide, the chest cavity may be opened with scissors, or the rodents 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 rodents, including the impact of euthanasia agents on tissues, please refer to the From:PHS POLICY ON HUMANE CARE AND USE OF LABORATORY ANIMALS CLARIFICATION REGARDING USE OF CARBON DIOXIDE FOR EUTHANASIA OF SMALL LABORATORY ANIMALS; Release Date: July 17, 2002; NOTICE: NOT-OD-02-062

Unintended recovery of animals after apparent death from CO2 (e.g., in necropsy coolers) is a documented occurrence. Institutions are reminded that such incidents constitute serious noncompliance with the PHS Policy and serious deviation from the provisions of the Guide for the Care and Use of Laboratory Animals (http://www.nap.edu/readingroom/books/labrats/). As such, the IACUC, through the Institutional Official, must promptly provide OLAW with a full explanation of the circumstances and actions taken. Prompt reporting in accordance with PHS Policy requirements (PHS Policy IV.F.3) is an essential component of the formal relationship between OLAW and PHS-Assured institutions."

Click here to see the entire source document on the OLAW website.

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
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, 1998.
2. Occupational Health and Safety in the Care and Use of Research Animals, National Research Council, 1997.

Texts:

1. Fox, Cohen, and Loew. Laboratry Animal Medicine. Academic Press, Inc., 1984.
2. Hawk and Leary, Formulary for Laboratory Animals. 2nd Edn., Iowa State University Press, 1999.
3. Hrapkiewicz, Medina, and Holmes, Clinical Laboratory Animal Medicine: An Introduction, 2nd Edn., Iowa State University Press, 1998.
4. Field and Sibold. The Laboratory Hamster & Gerbil. CRC Press, 1999.
5. Wolfensohn and Lloyd, Handbook of Laboratory Animal Management and Welfare. 2nd Edn., Blackwell Science. 1998.