Transcript The Laboratory Rat - The University of Tennessee College of

BIOMETHODOLOGY OF THE RAT
Office of Laboratory Animal Care
University of Tennessee, Knoxville
General Behavior
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Nocturnal
Photoperiod 12:12 or 14:10 for
breeding colonies
Non-aggressive, inquisitive,
trainable
Coprophagic
Accept single housing
Males unlikely to fight when
housed together
Body Weight Gain Chart
CD Rat
Reproduction
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Estrus Cycle 4–5 d
Gestation 21–23 d
Litter size 3-18 pups
Haircoat 8-9 d
Eyes open 10-14 d
Weaning 21 d
Postpartum estrus
Sexing
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Anogenital distance is longer in the
male and shorter in the female
Male
Female
Genital
Opening
Anus
Housing
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Adequate housing should provide the
following:
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Behavior and physiological needs
Social interaction
Clean, dry and safe area with adequate
ventilation, food and water
Visualization by personnel
Sufficient space to turn around and make
normal postural movements
Housing Recommendations
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Primary enclosure space recommendations per the
Guide for the Care and Use of Laboratory Animals
Rat
Weight
(g)
Floor Area
(in2)
Height
(in)
<100
17
7
Up to 200
23
7
Up to 300
29
7
Up to 400
40
7
Up to 500
60
7
>500
>70
7
Primary Enclosure
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Enclosure
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Locking wire bar lid
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Easy visualization
Solid bottom flooring
Bedding and enrichment
Water bottle
Feed
Microisolator top
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Reduces spread of
pathogens
Environmental Conditions
Room Recommendations
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Temperature 68 – 79⁰F
Humidity 30 -70%
Ventilation 10 -15 air changes/hr.
Light levels 130 -325 lux
Noises sustaining ≥85 db can cause
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Stress
Metabolic changes
Reduced fertility
Identification
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Temporary
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Markers and Dyes
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Hair Clipping
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Ear Tags
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Microchip
Permanent
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Ear Punches
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Tattoos
Identification
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Cage Card
Manual Restraint
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Use gloves
Grasp the tail 1-2
cm from the base
and place rat on
wire bar lid
Manual Restraint
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Palm the rat over
the back
Push the rat against
the wire bar
Advance hand
toward rat’s head
Manual Restraint
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Place index and
middle finger
around the neck
Keep the head
between the
middle and index
fingers
Manual Restraint
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The thumb and
second finger
should be placed
under opposite
axilla
The tail can be
held by the
opposite hand
Manual Restraint
Click to Watch Video
Mechanical Restraint
Plastic adjustable
restrainers
Decapicones
Blood Collection
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Survival Procedures
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Tail vein
Orbital plexus
Non-Survival Procedures
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Cardiac puncture
Axillary cut down
Cranial vena cava puncture
Blood Collection
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Blood Collection Guidelines
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Single blood draw
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≤1.0 ml per 100 grams of body weight
Multiple blood draws
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Maximum of 1.5 ml per 100 grams of body weight
within a 2 week period
Blood Collection
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Tail Vein
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Use for collecting
0.1–2.0 ml of blood
Heat source to dilate
blood vessel
Proper restraint
≤ 21 gauge needle
Blood Collection
Click to Watch Video
Blood Collection
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Orbital plexus
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Use for collecting up to
4 ml of blood
Anesthetize rat
Hold the head steady
Insert pipette in the
medial canthus of the eye
Rotate the tube between
thumb and finger
Keeping eyelids closed,
apply direct pressure
using gauze for hemostasis
Blood Collection
Click to Watch Video
Blood Collection
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Cardiac
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Use for collecting up
to 15 ml of blood
Anesthetize rat
≤21 gauge needle
Insert needle under
sternum at a 20⁰ angle
Aspirate slowly
Euthanize rat
Blood Collection
Click to Watch Video
Tissue Collection
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Tail Biopsy
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Limited to a maximum of 2 times
Maximum biopsy size of 5 mm
Analgesia/Anesthesia is required for rats 21 days
of age and older and in rats weighing 50 grams
or more
Compound Administration
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21 to 25 gauge needle
Maximum Administration Volumes (in ml/kg)
IM
0.1–0.2
IP
10–20
IV
PO
(bolus)
(gastric gavage)
5
10–40
SC
2-10
Compound Administration
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Subcutaneous (SC or SQ)
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Locate site of injection
Insert needle underneath
skin
Aspirate negative
pressure
Inject compound and
watch for SQ bleb
Pause before retracting
needle
Compound Administration
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Intraperitoneal (IP)
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Assistance is helpful
Locate lower right quadrant for injection site
Aspirate
If an unintended subcutaneous bleb occurs,
reposition the needle
Compound Administration
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Intravenous (IV)
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Lateral tail vein
Proper restraint
Heat source to dilate
blood vessel
After removing needle,
apply direct pressure
for hemostasis
Compound Administration
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Gastric Gavage (PO)
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Use a bulb-tipped gastric gavage needle
Measure length of needle from mouth to last
rib
DO NOT FORCE the needle down the
esophagus
Inject solution
Observe rat for signs of distress
Anesthesia
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When planning any procedure involving
anesthesia and/or surgery, please consult one
of the Laboratory Animal Veterinarians in the
Office of Laboratory Animal Care (OLAC) at
974-5634.
The Veterinarian can provide guidance and
detailed information in selecting the most
appropriate anesthetic and analgesic protocol
for your rat and procedure.
Aseptic Technique
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Surgical Prep
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After induction of anesthesia, clip hair from the
surgical site
Prep skin with povidone iodine, chlorhexidine or
other appropriate skin antiseptic
Scrub in a circular pattern, beginning in the center
and spiraling outward
Follow povidone iodine scrub with a 70% alcohol prep
Repeat Twice
End procedure with a light coat of povidone iodine
solution (not scrub) to the surgical site
Aseptic Technique
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Place a sterile drape
over the rat
Anything that touches
the surgical site must
be sterile
Non-absorbable
sutures/clips should be
removed in 7-14 days
Surgical Monitoring
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Prevent pain, hypoxia, and hypothermia
Monitor withdrawal reflex
Provide a source of external heat
Provide appropriate analgesics for postoperative pain management
Surgical Record
Euthanasia
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Inhalant Anesthetic Overdose
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Carbon Dioxide
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Isoflurane
End the procedure with a thoracotomy or cervical dislocation
Place rat in the chamber
Turn on CO2 flow into the chamber
Once the rat has stopped breathing, wait at least 1 minute before
removing the rat from the chamber
End the procedure with a thoracotomy or cervical dislocation
Substantially prolonged in neonates
Cervical Dislocation
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Performed on anesthetized rats weighing <200 grams
Must be performed by skilled personnel
Prevention of Infectious Disease
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Colony Health Surveillance
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Sentinel rats are tested on a routine basis
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Gross necropsy
Serological assays
Tests for parasites
Rat Antibody Production Test (RAP Test) and
PCR
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A test for cell lines and tumors for rodent viruses
Included in every animal use protocol
Occupational Health and Safety
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PPE (Personal Protective Equipment)
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Protects handler and rat
May include: gloves, shoe covers,
gowns, lab coats, masks, and head
covers
Zoonoses prevention
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Proper handling
Health surveillance programs
Routine sanitation
Proper use of PPE
Health Surveillance
Rat Serology (RADIL)
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Rat coronavirus (RCV)
Generic parvovirus (NS1)
Rat parvovirus (RPV)
Rat minute virus (RMV)
Kilham rat virus (KRV)
Toolan's H-1 (H-1)
Rat theilovirus (RTV)
Sendai virus
Pneumonia virus of mice
(PVM)
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Mycoplasma pulmonis
Reovirus 3
Lymphocytic choriomeningitis
virus (LCMV)
Cilia associated respiratory
bacillus (CARB)
Hantaan virus
Tyzzer’s
Mouse adenovirus (MAD1)
Procurement of Rats
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APPROVED vendors include Charles River,
Jackson Labs, Harlan, NCI-Frederick, and
Taconic
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Rats that are shipped from a non-approved
vendor source must be approved by OLAC before
the rats are procured.
An animal requisition form must be submitted
to the facility manager:
http://www.vet.utk.edu/olac/pdf/animal_acquisition_form.pdf
Quarantine
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Quarantine is required if receiving rats from
an unapproved vendor
The minimum quarantine period is six (6)
weeks
No experimental manipulations or breeding
can be initiated during the quarantine period
unless approval has been granted by an OLAC
veterinarian
Health Concerns
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Whenever possible,
pain and distress
should be eliminated
General appearance
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Lethargy
Aggressiveness
Hunched posture
Health Concerns
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Eyes
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Porphyrin staining = Stress
Teeth
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Overgrowth
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Tooth breakage
Malocclusion
Health Concerns
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Body Condition
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Swellings or Tumors
Poor Haircoat
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Ectoparasites
Stress
Cold
Poor Nutrition
Body Condition Scoring
Hickman DL, Swan M 2010
Reporting Signs of Pain or Distress
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For Rats That Require Veterinary Care
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Complete the red “Sick Animal”
cage card
Attach card to cage
Notify facility manager or
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Print Clinical Case Request Form
http://www.vet.utk.edu/olac/pdf/CLINICAL_CASE_REQUEST.pdf
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Fax form to the OLAC office 974-5649
Assessment of the animal’s condition and
treatments will be recorded on the “Sick
Animal” card
References
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Guide for Care and Use of Laboratory Animals. National Academy
Press, 2011.
Hickman, DL, Melissa Swan. 2010. Use of Body Condition Score
Technique to Assess Health Status in a Rat Model of Polycystic
Kidney Disease. J Am Assoc Lab Anim Sci 49:155-159.
Lawson, PT. Assistant Laboratory Animal Technician (ALAT)
Training Manual. The American Association for Laboratory Animal
Science, Memphis, TN. 2005.
Sharp, Patrick and Marie LaRegina. The Laboratory Rat. CRC
Press, Boca Raton, FL. 1988.
Presentation: Chris Carter, OLAC UTK