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The University of New Mexico


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Title: The University of New Mexico

The University of New Mexico The Office of
Animal Care and Compliance
  • present
  • Basic Biomethodology for Laboratory Rats
  • A learning module developed and presented by the

  • This module was developed to assist you in
    becoming proficient in performing common
    techniques in the rat

General Information
  • We all have an ethical responsibility to animals
    in terms of minimizing pain and distress
  • This can be accomplished, in part, by using
    proper animal handling and experimental
  • The PHS requires that animal care and use is
    based on the Guide for the Care and Use of
    Laboratory Animals
  • Personnel caring for animals should be
    appropriately trained

Entrance Procedures
  • Access to most UNM animal facilities is by
  • Sharing of card-keys could result in termination
    of your access privileges
  • Animal users are not provided with access to the
    animal facility until all training requirements
    are met

Personal Protective Equipment
  • All animal facilities require some level of
    protective clothing in order to protect the
    animals housed within from contaminants that may
    enter the facility via the personnel and to
    protect the personnel from exposure to animal
    allergens or other potential hazards

Examples of Protective Clothing
  • Lab Coats
  • Jumpsuits
  • Shoe Covers
  • Hair Bonnets
  • Masks
  • Gloves

Animal Records
  • All animals used in research require some kind of
    record and identification, even if it is only a
    cage card
  • Cage cards should include protocol number,
    investigator, species, strain, age/wt, sex,
    vendor, and date received

Microisolator Cage Technique
  • Some facilities require that all animal cages be
    opened inside a biosafety cabinet
  • The cages are sprayed with a disinfectant prior
    to placing them in the cabinet and prior to
    removing them from the cabinet
  • Hands are sprayed with disinfectant prior to
    handling the cage and again prior to handling the
    cage contents

Microisolator Cage Technique
  • Sometimes you must manipulate the cages on a cart
    or bench top
  • The cages and gloved hands are sprayed with a
  • The microisolator lid is removed and placed
    inverted on the cart or bench top
  • Hands are sprayed with disinfectant again
  • The wire bar lid is removed and placed on the
    inverted microisolator top

Assessing the General Health of Rats
  • A brief assessment of the health of every animal
    should be conducted prior to performing any
    technical procedures
  • The animal should be observed for signs of
    illness including
  • Ocular or nasal discharge
  • Rough hair coat
  • Abnormal posture
  • Uterine, rectal or penile prolapse
  • Limb abnormalities
  • Malocclusion
  • Dehydration
  • Dystocia
  • Abnormal behavior
  • Example of ulcerative podododermatitis
  • Example of Malocclusion

Assessing the General Health of Rats
  • Any signs of pain or distress should be reported
  • Observe the feed and water supplies to ensure
    that there is evidence that the animal has been
    eating and drinking
  • Group-housed animals will often fight, they
    should be observed closely for fight wounds and
    separated immediately if fighting is noted
  • Barbering may also be seen in group-housed rats
    of both sexes
  • The muzzle and other areas of the body are shaved
    by the dominant rat in the cage

Sexing rats
Notice the greater distance in the male
Notice the more prominent hairless strip in the
  • It is important that every animal handler be
    properly trained to distinguish between male and
    female rats
  • The anogenital space is almost twice as long in
    the male as it is in the female
  • It is difficult to differentiate the sex of
    neonates it is sometimes helpful to compare two
    animals side by side

Rat Behavior
  • Rats are nocturnal
  • Rats exhibit strong burrowing, gnawing and
    nesting behavior
  • Rats should be provided with bedding materials
    that encourage this behavior
  • Other environmental enrichment devices should
    also be used as appropriate, but must be approved
    by the Attending Veterinarian prior to use

  • When attempting to restrain rats, sudden, jerky
    moves should be avoided to decrease the
    likelihood of being bitten
  • Approaching with gentle confidence is best
  • Select the appropriate method of restraint for
    the procedure you wish to perform
  • Restraint by the tail is intended for short-term
    manipulations such as cage transfer

Tail Restraint
  • Notice how close to the base of the tail the
    handler is
  • Rats may be picked up by grasping the base of the
  • Do not grasp the tip of the tail as this may
    cause the skin to be stripped off
  • Never suspend a rat for long periods of time by
    its tail
  • Notice how the handler is supporting the weight
    of the rat

Two-Handed Restraint
  • Using two-handed restraint or mechanical devices
    is suggested for procedures requiring more than
    momentary restraint
  • Restraining the rat by the two-handed method will
    allow you to perform many technical procedures
    such as examination, injection and blood
  • There is a scruff and a thoracic variation of
    this technique

Scruff Restraint
  • Restrain the rat by grasping it at the base of
    the tail and placing it on a table or cart
  • While grasping the tail with one hand, grasp the
    nape or scruff of the neck with the other
  • Be sure to grasp enough skin so that the animal
    cannot turn its head to bite you

Thoracic Restraint
  • Restrain the rat by grasping it at the base of
    the tail and placing it on a table or cart
  • While grasping the tail with one hand, grasp the
    rat around the thoracic area with the other do
    not squeeze too tight or you could cause
    breathing difficulties
  • Be sure to keep the animals forelimbs above your
    thumb and index finger as this will protect you
    from being bitten

Thoracic Restraint - Variation
  • A variation of this method places the animals
    head between the first and second fingers while
    the forelimbs are held between the thumb and
    forefinger, and the third and fourth fingers
  • Again, care must be taken to hold the animal
    tight enough to prevent it from biting you, yet
    loose enough to prevent breathing difficulties

  • Monitor the condition of the animal the entire
    time it is restrained
  • Observe the breathing rate and color of the ears,
    nose and oral cavity
  • Release the animal immediately if there are any
    signs of gasping or change in coloring from pink
    to blue

Mechanical Restrainers
  • Plexiglass restrainers are available from
    different manufacturers in a variety of styles
  • They allow the user to have both hands free for
  • Depending upon the type of device, the animal can
    be placed in the restrainer either head or tail
  • Cage tops can also be used to restrain rats for
    procedures on the tail

Head-First Restrainers
  • Grasp the rat by the base of the tail with one
    hand and cover the restrainer with the other
  • Most rats, once they are shown the entrance to
    the tunnel, will readily enter the restrainer

Head-First Restrainers
  • If the restrainer has a securing device, affix it
    firmly to prevent the rat from exiting the

Tail-First Restrainers
  • These are appropriate for such procedures as
    tail-vein injections and blood collections
  • Grasp the rat by the base of the tail and slide
    its hindquarters first into the restrainer
  • Use the slot as a tail guide

Tail-First Restrainers
  • Once the rat if fully in the restrainer, insert
    the securing device to prevent the animal from
    exiting the apparatus

Identification Methods
  • It is important to select the appropriate
    identification method for your research purposes
  • This should be based upon the age of the animal,
    the number of characters you wish to include, and
    the duration of the experiment
  • You should record the identification information
    on the cage card in the event that clarification
    of the numbers or characters becomes necessary
    for any reason

Identification Methods
  • Indelible markers can be used for short-term
  • Ear-punches, microchips, and tattooing are all
    permanent procedures
  • Ear tags can be long-term, but there is always a
    chance they will become detached from the ear

Temporary Identification
  • Non-toxic, permanent markers can be used to
    temporarily mark the fur, tail, or skin of the
  • This ink usually lasts 3-4 days without the need
    to remark

Ear Punches
  • Different types of ear punches are available
  • Ear punches should be sterile prior to use
  • Extra ear punches should always be available as
    they become dull with repeated use

Ear Punches
  • Restrain the rat by the scruff using the method
    described previously
  • The punch should be placed approximately 3mm from
    the edge of the ear pinna
  • If the punch is placed to close to the edge it is
    likely to tear and become difficult to read
  • You should also take care not to place the punch
    too far towards the inside of the ear to avoid
    injuring the animal
  • The tissue removed can also be used for genotyping

Ear Punches
  • Sanitize the ear punch between each cage of
    animals with 70 ethanol
  • Chlorinated compounds will cause the punch to
    become corroded

Microchip Transponders
  • Often called Pit Tagging, they are implanted
    subcutaneously between the scapulae for permanent
    identification of individual animals
  • Each microchip is encrypted with a unique,
    non-replicable number and are read with a
    portable, hand-held scanner

Microchip Transponder
  • To implant these chips, the rat must be briefly
  • The hair is removed from the insertion site by
  • The area is prepped with an iodophor, followed by
  • The implantation needle, with the syringe
    attached, is purchased in a sterile package
  • Make a tent from the loose skin at the implant

Microchip Transponders
  • Insert the needle subcutaneously, with the bevel
    up, and depress the plunger

Note that the bevel is facing up
  • Once the needle is removed the injection site
    should be observed for bleeding
  • If bleeding is noted, apply pressure with a
    sterile gauze pad, or a drop of super glue to the
    entry site

  • Tattooing can be used on both neonates and adults
    as a permanent method of identification
  • Anesthesia is not required, but can be used to
    immobilize the animal
  • A tattoo device can be used to write numbers or
    other characters on the tails of adult mice
  • It can also be used to tattoo the footpads of
    both neonatal and adult mice

  • The use of tattoo equipment requires further
    hands-on training
  • All tattoo equipment should be disinfected prior
    to use and sterilized between animal rooms

Ear Tags
  • Ear tags are another means for identifying rats
  • Ear tags can be imprinted by the manufacturer
    with several digits or letters
  • Special attention should be given to the
    placement of the ear tag
  • Improperly placed tags can become easily detached
    from the ear, torn out when the rats fight, or
    inadvertently become caught in the wire bar lid

Ear Tags
  • Restrain the rat as previously described
  • Place a sterile ear tag into a sanitized ear tag

Ear Tags
  • Locate the proper position for placement (numbers
  • Apply the tag to the base of the ear,
    approximately 3mm from the edge of the pinna
  • Do not apply the tag too close to the center of
    the ear as this may cause inflammation,
    necessitating removal of the tag
  • Do not apply the tag too close to the outer edge
    of the pinna as this may cause the tag to become
    entangled with the foot of the animal or in the
    wire bar lid, causing it to become detached from
    the ear

Toe Clipping
  • Toe clipping for identification is not
  • Toe clipping requires scientific justification
    and approval of the IACUC
  • To clipping requires specific training in the
    procedure by Animal Facility Staff

Genotyping Tail Snips
  • Most commonly, genotyping of rats is accomplished
    by amputating a small portion of the distal tail
  • This is best performed when the pup is 10 15
    days old
  • 5.0 mm or less of the distal tail is removed for
    this procedure
  • The tail should be sprayed with a topical
    hypothermic, or the animal may be anesthetized

Genotyping Tail Snips
  • A scalpel or scissors can be used to remove the
  • The instruments are steril at the beginning of
    the procedure and sanitized between animals
  • You must assure that adequate hemostasis has been
    achieved before returning the animal to its cage
  • Surgical glue, silver nitrate, or direct pressure
    with a sterile pad can be used for this purpose
  • Note the tissue removed during ear punching can
    be used for rapid screening of rats
  • You may also be able to use blood, hair, saliva,
    or feces for genotyping consult with the

  • Various routes exist for injecting rats
  • Discuss the appropriate route, volume, site and
    needle selection with the veterinarian
  • All injections must be described in your approved
  • All injections must be performed using sterile
    needles and syringes
  • A new needle and syringe should be used for each
    cage of rats

Intramuscular Injections - IM
  • Regardless of the method used for IM injections,
    it must be noted that the sciatic nerve runs
    along the length of the femur
  • It is very important to avoid injuring this nerve
  • This is best accomplished by pointing the needle
    caudally rather than cranially, however, the
    quadriceps muscle can be used by an experienced
  • It is imperative that the rat is properly
    restrained either two handed with an assistant
    injecting the animal, or using anesthesia
  • Swab the area with 70 ethanol before placing the
    needle and aspirate to look for blood before

Intramuscular Injections - IM
  • Location of the sciatic nerve and target
    injection site

Intramuscular Injections - IM
  • Have one person restrain the rat as described
  • The second person will secure the rear foot
    nearest to the first persons lower thumb
  • Swab the area with 70 ethanol
  • Insert the needle, bevel up, into the caudal
    thigh at a 45 angle
  • Aspirate to ensure that you have not entered a
    blood vessel then slowly inject the material

Intramuscular Injections - IM
  • You can perform the injection without anesthesia
    but this is not recommended
  • Two people must be used for IM injections on
    unanesthetized animals
  • One person restrains the rat, while the second
    person performs the injection
  • The quadriceps can be used, but the caudal thigh
    is recommended

Subcutaneous Injections SC or SQ
  • Restrain the rat with anesthesia or as described
  • A second person will use their thumb and
    forefinger to make a tent of skin over the scruff
  • Prep the area with 70 ethanol
  • Insert the needle, bevel up, at the base of the
  • The needle should be inserted parallel to the
    skin and should be directed toward the posterior
    of the animal
  • Aspirate to ensure proper placement and inject
    the material

Intraperitoneal Injections IP
  • Restrain the rat with anesthesia (preferred), or
    by the scruff method or two-handed method using
    two persons
  • Expose the ventral side of the animal
  • Prep the site with 70 ethanol
  • The sterile needle should be placed, bevel up, in
    the lower right or left quadrant of the animals
  • Insert the needle at a 30 angle
  • Aspirate and inject the material

Anesthetized Animal
Intravenous Injections IV
  • Warm the rats tail in a bowl of warm water, or
    under a heat lamp or other heating device
  • Be sure not to OVERHEAT the animal
  • The temperature should not exceed 85 - 90 F at
    the level of the animal
  • Remove the rat from the heat source immediately
    should any change in respiration rate or
    excessive salivation be observed

Intravenous Injections IV
  • Place the animal in a restraint device and
    stabilize the tail between the thumb and
    forefinger of the hand that will not be
    manipulating the syringe
  • Or restrain the animal with an anesthetic and
    stabilize the tail between the thumb and
    forefinger of the hand that will not be
    manipulating the syringe

Intravenous Injections IV
  • Prep the tail with 70 ethanol
  • Attempt the injection starting at the middle or
    slightly distal part of the tail
  • With the tail under tension insert the needle,
    bevel up, approximately parallel to the vein and
    insert the needle at lest 3 mm into the vein
  • DO NOT ASPIRATE this will cause the vein to
  • Inject the material in a slow, fluid motion
  • You should see the vein blanch if the needle is
    properly positioned
  • If any swelling at the injection site or
    resistance to injection occurs, remove the needle
    and reinsert it slightly above the initial site

Intradermal Injections ID
  • In order to perform ID injections the animal
    should be anesthetized
  • Shave an injection site on the back of the animal
    to remove the hair
  • Swab the site with 70 ethanol
  • Insert the needle into the skin, bevel up,
    holding the needle nearly parallel to the plane
    of the skin
  • Do not aspirate
  • Inject the material
  • The volume of the injection should be limited to
    50 µl per site to avoid tissue trauma
  • A properly performed ID injection should result
    in a small, round skin welt

Injection Sites and Volumes
  • SQ in the Scruff Maximum 10 ml 20-25ga
  • IM Caudal Thigh 0.3 ml 21ga needle
  • IP Lower Ventral Quadrants 10 ml 20-25ga
  • ID Lateral Abdomen/Thorax 0.05 ml 25-27ga
  • IV Lateral Tail Vein 0.5 ml 20-25ga needle

Oral Gavage
  • Gavaging is used to dose an animal with a
    specified volume of material directly into its
  • Only a specialized, commercially available gavage
    needle should be used for this procedure

Oral Gavage
  • Fill the syringe with the appropriate volume of
    material and attach the needle
  • Restrain the animal by the scruff
  • Place the tip or ball of the needle into the
    animals mouth

Make sure you measure the gavage needle for
proper length
Oral Gavage
  • Slide the tip gently past the back of the tongue
  • The needle should slide easily down the esophagus
    if properly placed
  • If any resistance is met, remove the needle and
  • Do not aspirate once the needle is properly
    placed administer the material

Blood Collection
  • It is important to select the proper method of
    blood collection that corresponds to the volume
    required for your research purposes
  • Some methods are intended for survival and others
    are not
  • Consult the veterinarian for more information

Typical Blood Collection Sites Includes the
  • Retro-orbital Sinus Blood Collection Survival
  • Submandibular Blood Collection Survival
  • Lateral Tail Vein Blood Collection Survival
  • Saphenous Vein - Survival
  • Intracardiac Puncture Blood Collection

Retro-Orbital Sinus
  • The retro-orbital sinus is the site located
    behind the eye at the medial or lateral canthus
  • This venous sinus is located just underneath the
    conjunctival membrane
  • No more than 2 of the blood volume should be
    removed at one sampling
  • The blood volume of a rat is approximately 5-7
    of body weight
  • A 250 gm rat has a circulating blood volume of
    about 15-35 ml, so no more than 500 µl of blood
    should be removed at one single bleeding

Retro-Orbital Sinus
  • Rats should not be bled more frequently than
    every 3 weeks unless smaller samples are
  • Animals should be anesthetized prior to
    performing this procedure
  • Inhalant anesthetic is the preferred method
  • It is imperative that the animal is properly
    restrained or severe injury to the eye or
    surrounding tissue could occur

Retro-Orbital Sinus
  • Restraining the animal by the scruff method and
    tightening up slightly to the loose skin around
    the neck will cause the eye to bulge slightly
  • Care should be given to ensure the animal does
    not have difficulty in breathing

Retro-Orbital Sinus
  • With a gentle rotating motion, insert the tube
    through the sinus membrane
  • Continue rotating the tube at the back of the
    orbit until blood flows

Retro-Orbital Sinus
  • Collect the appropriate volume of blood
  • Ensure good hemostasis with a clean gauze pad
    before returning the animal to its cage
  • To become proficient at this technique,
    additional training outside the scope of this
    text is required
  • Please contact the ARF for appropriate training

  • Veins draining the eye and submandibular area
    meet at the rear end of the cheek pouch
  • This provides a convenient and consistent source
    of blood
  • Prepare the animal as outlined earlier for
    retro-orbital blood collection

  • Using a 25ga needle, nick the submandibular vein
  • Allow the blood to drip into a collection device
  • Once the sample is collected, assure proper

Lateral Tail Vein
  • Tail nicking is a survival procedure that can be
    used to collect up to 500 µl of blood from the
    lateral tail veins
  • This method must be used with caution, as when
    improperly performed, permanent tail injury or
    amputation may occur
  • Prepare the animal as outlined earlier for tail
    vein injections

Lateral Tail Vein
  • Using a 11 scalpel blade, gently nick the
    lateral tail vein in the general area around the
    midline of the tail
  • Start at least halfway down the tail so if there
    is a problem, you can nick the tail above the
    initial site and still obtain your blood sample
  • Allow the blood to flow into an appropriate
  • Do not attempt to squeeze the tail or milk blood
    from the tail this may cause tissue damage and
    contamination of the blood sample
  • When the sample is collected, ensure good
    hemostasis with a sterile gauze pad, surgical
    glue, or silver nitrate

Saphenous Vein
  • The saphenous vein may also be used for blood
  • Restrain and extend the hind leg applying gentle
    downward pressure above the knee joint.
  • Wipe the shaved area with alcohol or sterile
    lubricating gel and use a 25-gauge needle to
    puncture the vein (The vein is next to the dark
    highlight in the picture below).
  • If done correctly a drop of blood forms
    immediately at the puncture site and can be
    collected in a micro-hematocrit tube.

Saphenous Vein
  • If collecting blood from the right leg, the fold
    of skin between the abdomen and cranial thigh
    surface is used to fix the leg
  • The hair is then removed from the outer surface
    of the fixed leg
  • The vein should now be visible on the surface of
    the thigh
  • Prep the area with 70 ethanol
  • A 25ga needle is held almost parallel to the
    saphenous vein the vessel is punctured it is
    not necessary to lance the vein
  • The appropriate capillary tube is held on a 45
    angle with one end of the tube at the edge of the
    drop of blood collecting on the leg surface

Saphenous Vein
  • Approximately 300 µl of blood can be collected
    from an adult rat using this method
  • Flex the rats foot to reduce the flow of blood
  • Slight pressure is then applied to the puncture
    site with a gauze compress until hemostasis occurs

Intracardiac Puncture
  • This procedure must be performed under deep
    anesthesia and is a NON-SURVIVAL procedure
  • Once the animal is anesthetized, prep the chest
    with 70 ethanol
  • Insert the needle at the base of the sternum,
    bevel up, into the thoracic cavity at a 15-20
    angle directed just to the left of midline
  • Aspirate slowly
  • If blood begins to flow into the syringe,
    continue to aspirate with steady, even pressure
  • If no blood is seen reposition the needle and try

Intracardiac Puncture
  • Once the required blood volume is collected, the
    rat is euthanized while still deeply anesthetized
  • Up to 10 milliliters or more of blood may be
    collected from an adult rat using this method

  • This module will provide a brief introduction to
    analgesia and anesthesia in the rat
  • Your veterinarian or ARF personnel should always
    be consulted for advice on selection and
    administration of analgesia or anesthesia
  • The use of analgesics and/or anesthetics must be
    described in your approved animal use protocol
  • There is a drug formulary on the OACC website
    that lists drugs, dosages, and uses for various

  • Injectables used for anesthesia and analgesia
  • Typically given IP or IM but may be given SQ
  • It is important to weigh the rat prior to dosing
    with an injectable anesthetic to avoid over or
    under dosing the animal

  • Topicals as an adjunct to, or in lieu of
    injectable analgesics, topical anesthetics may
    also be used
  • These long-acting agents are painted or dropped
    into the surgical wound before the skin is closed
  • To facilitate retro-orbital sinus blood
    collection, an opthalmic anesthetic is used as a
    topical analgesic

  • Inhalants the most commonly used inhalant at
    UNM is isoflurane
  • Isoflurane is administered in 10002 - induction
    concentrations of isoflurane are 3-4 and
    maintenance concentrations are 1.25-1.75
  • Inhalant anesthetics must be used with a
    scavenging device
  • Contact the veterinarian for further training in
    the appropriate use of the anesthesia machines
    available at UNM

  • Anesthetized animals must be monitored closely
    during the procedure to assure that they are
    maintained in the proper anesthetic plane
  • If the plane is too light the animals may move or
  • If the plane is too deep the animals may die
  • The plane can be assessed by pinching the toe,
    tail, or ear of the animals
  • Any reaction from the animal indicates that the
    anesthesia is too light and additional anesthesia
    should be given

  • The respiration and color of the mucous membranes
    and exposed tissue of the animal should also be
    closely monitored
  • The respiration rate should be even
  • An increase in respiration indicates that the
    anesthesia is too light
  • A deep, shallow, decreased or irregular
    respiration indicates that the anesthesia is too

  • The color of the mucous membranes and exposed
    tissues should be bright pink to red
  • Dusky grey or blue color is indicative of an
    anesthetic plane that is too deep
  • Core body temperature can also be monitored in
    rodents the most common anesthetic complication
    is hypothermia
  • Measures must be taken to control the body
    temperature during and after anesthesia

  • Place the animal on a clean, dry gauze or paper
    towel to avoid contact with the bedding which
    may be inadvertently inhaled and result in
  • Once the animal has reached sternal recumbency
    and appears to making a normal recovery, it may
    be returned to the animal holding area
  • Animals should be watched for several days
    following a procedure

  • The definition of euthanasia is the intentional
    induction of a painless death
  • The veterinarian should always be consulted for
    advice on selection and administration of
    euthanasia agents
  • The euthanasia method must be fully described in
    your approved animal use protocol

Euthanasia C02
  • Compressed carbon dioxide gas is the only
    recommended source of C02 for euthanasia
  • Carbon dioxide generated from dry ice is NOT
  • With an animal in a chamber, an optimal flow rate
    should displace 10-20 of the chamber volume per
    minute until the mouse is unconscious

This flow rate is associated with a rapid loss of
consciousness and minimal distress to the animal
Euthanasia C02
  • Once the animal is unconscious the flow rate can
    be decreased
  • Gas flow should be maintained for at least 1
    minute following apparent clinical death
  • Death should be verified by the absence of the
    heartbeat, performing cervical dislocation, or
    perforating the diaphragm prior to disposal of
    the animal

Euthanasia Injectable Inhalant
  • Injectable anesthetics can also be used for
    euthanasia when administered at higher doses
  • Barbituate anesthetics produce rapid and humane
    euthanasia when injected IP
  • Halothane is the most effective inhalant
    anesthetic for euthanasia, but isoflurane can
    also be used
  • Inhalants are best utilized with the open drop
    method using a closed receptacle containing
    cotton or a gauze soaked with the liquid
  • You must prevent direct contact of the animal
    with the liquid anesthetic

Euthanasia Physical
  • Cervical dislocation or decapitation, when
    properly performed, is a humane method of
  • Cervical dislocation can only be performed on
    small rats (lt125gms)
  • Animals MUST be anesthetized prior to cervical
    dislocation or decapitation unless scientifically
    justified and approved by the IACUC
  • Fetuses and neonates are resistant to many
    methods of euthanasia and special considerations
    must be given to this age group

This Concludes Module 4 Basic Biomethodology
for Laboratory Mice
  • Please download the exam, complete it, then
    e-mail it to
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