The University of New Mexico

1
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
  OACC

2
Introduction

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

3
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
  techniques
• 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

4
Entrance Procedures

• Access to most UNM animal facilities is by
  card-key
• 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

5
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

6
Examples of Protective Clothing

• Lab Coats
• Jumpsuits
• Shoe Covers
• Hair Bonnets
• Masks
• Gloves

7
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

8
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

9
Microisolator Cage Technique

• Sometimes you must manipulate the cages on a cart
  or bench top
• The cages and gloved hands are sprayed with a
  disinfectant
• 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

10
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

11
Assessing the General Health of Rats

• Any signs of pain or distress should be reported
  immediately
• 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

12
Sexing rats
Notice the greater distance in the male
Notice the more prominent hairless strip in the
female

• 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

13
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

14
Restraint

• 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

15
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
  tail
• 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

16
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
  collection
• There is a scruff and a thoracic variation of
  this technique

17
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

18
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

19
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

20
Precautions

• 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

21
Mechanical Restrainers

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

22
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

23
Head-First Restrainers

• If the restrainer has a securing device, affix it
  firmly to prevent the rat from exiting the
  apparatus

24
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

25
Tail-First Restrainers

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

26
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

27
Identification Methods

• Indelible markers can be used for short-term
  identification
• 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

28
Temporary Identification

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

29
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

30
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

31
Ear Punches

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

32
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

33
Microchip Transponder

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

34
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

35
Tattooing

• 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

36
Tattooing

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

37
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

38
Ear Tags

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

39
Ear Tags

• Locate the proper position for placement (numbers
  up)
• 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

40
Toe Clipping

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

41
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

42
Genotyping Tail Snips

• A scalpel or scissors can be used to remove the
  tissue
• 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
  veterinarian

43
Injections

• 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
  protocol
• All injections must be performed using sterile
  needles and syringes
• A new needle and syringe should be used for each
  cage of rats

44
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
  person
• 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
  injecting

45
Intramuscular Injections - IM

• Location of the sciatic nerve and target
  injection site

46
Intramuscular Injections - IM

• Have one person restrain the rat as described
  earlier
• 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

47
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

48
Subcutaneous Injections SC or SQ

• Restrain the rat with anesthesia or as described
  earlier
• 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
  tent
• 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

49
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
  abdomen
• Insert the needle at a 30 angle
• Aspirate and inject the material

Anesthetized Animal
50
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

51
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

52
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
  collapse
• 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

53
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

54
Injection Sites and Volumes

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

55
Oral Gavage

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

56
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
57
Oral Gavage

• Slide the tip gently past the back of the tongue
• The needle should slide easily down the esophagus
  if properly placed
• DO NOT FORCE!!!
• If any resistance is met, remove the needle and
  reinsert
• Do not aspirate once the needle is properly
  placed administer the material

58
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
Following

• Retro-orbital Sinus Blood Collection Survival
• Submandibular Blood Collection Survival
• Lateral Tail Vein Blood Collection Survival
• Saphenous Vein - Survival
• Intracardiac Puncture Blood Collection
  Non-Survival

59
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

60
Retro-Orbital Sinus

• Rats should not be bled more frequently than
  every 3 weeks unless smaller samples are
  collected
• 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

61
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

62
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

63
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

64
Submandibular

• 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

65
Submandibular

• Using a 25ga needle, nick the submandibular vein

• Allow the blood to drip into a collection device

• Once the sample is collected, assure proper
  hemostasis

66
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

67
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
  receptacle
• 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

68
Saphenous Vein

• The saphenous vein may also be used for blood
  collection
• 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.

69
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

70
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

71
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
  again

72
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

73
Anesthesia/Analgesia

• 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
  species

74
Anesthesia/Analgesia

• 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

75
Anesthesia/Analgesia

• 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

76
Anesthesia/Analgesia

• 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

77
Monitoring

• 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
  struggle
• 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

78
Monitoring

• 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
  deep

79
Monitoring

• 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

80
Recovery

• 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
  asphyxiation
• 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

81
Euthanasia

• 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

82
Euthanasia C02

• Compressed carbon dioxide gas is the only
  recommended source of C02 for euthanasia
• Carbon dioxide generated from dry ice is NOT
  acceptable
• 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
83
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

84
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

85
Euthanasia Physical

• Cervical dislocation or decapitation, when
  properly performed, is a humane method of
  euthanasia
• 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

86
This Concludes Module 4 Basic Biomethodology
for Laboratory Mice

• Please download the exam, complete it, then
  e-mail it to KMirowsky_at_salud.unm.edu