Basic Technique for
Small Laboratory Animal
Aunchalee Sirimontaporn
National Laboratory Animal Center
Basic Technique for Mouse & Rat
1.
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8.
Handling /Restrain
Sexing
Identification
Administration
Blood Collection
Anesthesia
Euthanasia
Necropsy
Handling /Restrain

Critical factor for research successful

Depend on species, size, behavior of animal

Purpose of experiment
Handling /Restrain
• Removing an animal from Cage
• Restraint for Lab Practice
• Restraining Devices
MOUSE
Mouse Handling
Removing a mouse from Cage
Hold the base of the tail
by hand grasping the base of
the tail between thumb and
index finger
or Use the rubber-tipped
forceps to prevent damage to
tail
Mouse Handling
a) Two hand method
Gentle pull the tail with dominant handed & scruff the loose skin
Mouse Handling
b) One hand method
Scruff the loose skin
Handling /Restrain
Restrainer
Mouse Restraint
Restraining Devices
Rat Handling
o Removing animal from Cage
o Restraint for Lab Practice
o Restraining Devices
Removing animal from Cage
Scruff the loose skin at the back
Grasping whole body method
Restraint for Lab Practice
Scruff the loose skin at the back
Grasping whole body method
Observe animal breathing
Restraining
Devices
Restraining
Devices
Newborn Retrieval from Cage
Should be carefully and not disturb mother to cause the
cannibalism or neglect
Sexing Rat / Mouse
Newborn
Anogenital space
(genital papilla and anus):
MALE > FEMALE
Male
Female
Sexing Mouse
Adult
It’s clear
can see
Testicles
Male
Female
Sexing Rat
Adult
It’s clear
can see
Testicles
Male
Female
Identification
o Rapid and easy to apply individual and give the large
of numbers for unique ID
o Easy to read
o Humane, and in compliance with regulatory
agencies for the care and use of laboratory animals
o Base on age, number of characters and duration of
all the time for experiment.
Identification
Temporary Methods
Permanent Method
oEar punch
oClipped/Shaved Fur
oWriting by non-toxic dye
oEar Tag
oTattoo
o Microchip
Identification
Used permanent marker to write
number, bar on the tail or ears
for short-term individual ID
Identification
Ear punch
Used the international Code
Identification
tattoo
Identification
Ear Tag
•
•
Tag size must be appropriately for the species/age
Does not cause ear irritation or trauma
Identification
Microchips transponder
Routes of administration
o Ensuring proper animal restraint
o Selecting the proper site or route by
• nature and physiological of agent
• animal species
• local anatomy of administrated site
• project purpose
o Selecting the proper size and length of needle by
• animal age
• Injection site
• Characteristic of substance
Routes of administration
o Topical: local effect, substance is applied directly where
action is desired such as epicutaneous, inhalation, eye
drops, ear drop
o Enteral: desired effect is systemic, substance is given via
the digestive tract by Oral or Gavage or Rectal
o Parenteral: desired effect is systemic, substance is given
by other routes than digestive tract such as venous
injection
Routes of administration
o Intragastic Feeding: though the mouth into the stomach
via tube
o Subcutaneous Injection (SC): under the skin
o Intradermal Injection (ID): into layer of skin
o Intramuscular injection (IM): into muscle
o Intraperitoneal injection (IP): into abdominal cavity
o Intraveneous injection (IV): directly into vascular system
Intragastic Feeding
The Gavage tube has a ball-like enlargement at the
tip to prevent injury
•
Before inserting the tube , line it up
the ball at the level of the xiphoid
cartilage (last rib) to determine how
far the tube inserted to stomach.
• Insert the tube 45o angle with
horizontal plane
• Do not force
Subcutaneous Injection
• Place in the vascular space
between the skin and
underlying muscle
• Slow absorption
• Make a tent or pocket by
lifting the skin
• The needle should move freely
under the skin and no blood
after aspirated
Subcutaneous Injection
Intradermal Injection
• Give into the thick layer of skin
(between epidermis and dermis
layer)
• Use needle size ≥ 25G
• The most frequently used site is
over the shaved back that can see
the bleb or pocket of fluid
Intradermal Injection
Intramuscular injection
• Into the muscle of upper thigh hind limbs
• Intermediate rate of absorption
• Very small volumes per site
• Aspirate before injected substance
Intramuscular injection
Intraperitoneal injection
• Directed into abdominal
cavity above the inguinal
region (Lower- right quarter)
• Avoid to penetrate into organ
• Aspirate prior injection
Intraveneous injection
• Use lateral tail veins that start
lower on tail in order to move up
when practice’s fail
• Should treated the injected site
with antiseptic before puncturing
• Not easy for beginner
(requires training and practice)
• Aspirate prior injection
(Lawson, 2000)
Recommended injection Volume and Injection Sites
Species
Intravenous
Mouse
Later tail vein
0.2 ml.< 25G
Rat
Later tail vein
0.5 ml.< 23G
Guinea pig
Ear vein
Sphenous vein
0.5 ml.< 23G
Rabbit
Marginal ear vein
1-5 ml.< 21G
(Lawson, 2000)
Intraperitoneal
Imtramuscular
Subcutaneous
2-3 ml. < 25G
Quadriceps
posterior high
0.05 ml.< 23G
Scruff back
2-3 ml. < 20G
5-10ml. < 21G
Quadriceps
posterior high
0.3 ml.< 21G
Scruff back
5-10ml. < 20G
10-15ml. < 21G
Quadriceps
posterior high
0.3 ml.< 21G
Scruff back
5-10ml. < 20G
50-100ml. < 20G
Quadriceps
posterior high
0.5 ml.< 20G
Scruff back
30-50ml. < 20G
Collection of Blood
• Purpose of blood collection
• Type of blood required (Arterial or veins)
• Volume of blood required
• Duration and frequency of sampling
• Health status of animal being bled
• Impact animal welfare
• Potential for stress-induce effect on biochemical and hematological
parameters
• Training and experience of the technique
(NCI Frederic ACUC, 2006)
Collection of Blood
 Blood Sampling
Total blood volume in laboratory animals (Diehl, et al.,2001)
Species
Mouse
Rat
Guinea pig
Rabbit
Percentage of total blood volume of Body Weight
(%)
Recommended Mean
Range of means
7.2
6.3-8.0
6.4
5.8-7.0
7.5
6.7-9.2
5.6
4.4-7.0
Collection of Blood
 Blood Sampling
Limit Volume and recovery periods
Single sampling
Multiple Sampling
% Circulatory
% Circulatory
Approximate
Approximate
Blood Volume
Blood Volume
recovery period
recovery period
removed
removed in 24 hr.
7.5%
10%
15%
1 week
2 weeks
3 weeks
≤ 1.0 %*
7.5%
10-15%
20%
24 hr.
1 week
2 weeks
3 weeks
*If frequent sample are necessary, the use of cannulation as a less stressful
alternative to repeated venepuncture should be considered. (NC3Rs, 2012)
Collection of Blood
 Blood Sampling
Total volume and relevant Collection Volumes
Total
Blood
Volume
7.5% of
Circulatory
Blood
Volume
10% of
Circulatory
Blood
Volume
15% of
Circulatory
Blood
Volume
20% of
Circulatory
Blood
Volume
Mouse (25 g)
1.8 (ml)
0.1 (ml)
0.2 (ml)
0.3 (ml)
0.4 (ml)
Rat (250 g)
16 (ml)
1.2 (ml)
1.6 (ml)
2.4 (ml)
3.2 (ml)
Guinea pig
(900 g)
62 (ml)
4.7 (ml)
6.2 (ml)
9.3 (ml)
12.4 (ml)
Rabbit (4 kg)
224 (ml)
17 (ml)
22 (ml)
34 (ml)
45 (ml)
Species
(Weight)
Sites of Blood Collection
oOrbital Sinus/Plexus puncture
o Venous(Venipuncture)
• Tail vein
• Spheneous vein
• Jugular Vein
• Superficial temporal vein (Facial vein)
• Posterior Venacava
o Cardiac Puncture
o Axiliary Vessels
Orbital Sinus/Plexus
o Carried out under general anesthesia
o Penetrating the Orbital Sinus/Plexus with a
glass capillary tube or Pasteur pipette
o Required high level skill and competence
Orbital Sinus/Plexus
o Insert the tip of capillary tube at medial cantus
o give gentle pressure and rotate through the sinus
membrane continue rotating until blood flows
o Use clean gauze pad to be hemostasis
Orbital Sinus
Tail vein
o Anesthesia may be not necessary, but animals
must be use restrainer properly
o Vasodilatation for promote bleeding (exposing to
40o C for 10 sec. )
o Do not squeeze or milking the blood that may be
cause tissue damage and contamination blood
with tissue fluid
Tail vein
Two method
o Direct: using butterfly needle or needle and
syringe
o Indirect: cutting off tip of tail
Tail vein
Direct: using butterfly needle
or needle and syringe
o Use butterfly needle puncture
of lateral tail vein
o Not require anesthesia,
vasodilatation for promote
bleeding
Tail vein
Indirect: cutting off tip of
tail
o Commonly used in mice
o Restricted to tail tip
0.5-1.0 mm. removed
o Not suitable for older animal
Saphenous vein
o Creating a tiny puncture in the
saphenous vein on lateral side
of the lower rear leg
o Collection of the blood into a
capillary tube
o Multiple samples can be
collected in the same day
Jugular vein
o Carried out under
general anesthesia
o Penetrating into the
bleeding Area (clavicle
bone)
Bleeding Area
Superficial temporal vein (Facial vein)
o Commonly used in mice
o Do not anesthesia
o Locate the hairless
freckle on the side of the
jaw.
o Puncture the freckle
with the lancet/ needle.
ตาแหน่ งเก็บเลือด
ขั้นตอนการเก็บเลือด
Cardiac Puncture
o Carried out under general
anesthesia
o Palpate the heart beats
o Insert needle from the left side or
Insert needle under the sternum
o Withdrawn the blood
o Only used for terminal
Axillary Vessels
o Carried out under general
anesthesia
o use the surgical scissor cut like
a cup of auxiliary
o cut vessels and collected blood
o contaminate of body fluid
o only used for terminal
Posterior vena cava
o Carried out under general
anesthesia
o Open abdominal by aseptic
technique
o Collected blood from
Posterior vena cava
o Only used for terminal
Yes
Anesthesia
No
Blood collection technique and Blood Collection Volume
Blood Collection Volume
Species
Route
Mouse
Rat Guinea Rabbit
(ml)
(ml) pig (ml) (ml)
Superficial temporal veins
Sapheneous/Pedal vein
Marginal ear
Lateral tail vein
Central ear artery
Jugular vein
Tail tip amputation (<1-3mm)
Retrobulbar sinus/plexus
Cardiac, Axillary region,
Posterior vena cava
0.1-0.2
-
-
0.15
0.2
0.1-0.3
-
-
-
0.5-1.0
0.05-0.2
0.1-2.0
-
-
-
-
-
0.5-10
0.05-0.2
0.1-2.0
-
-
0.01
0.2
0.1
0.5-2.0 1.0-4.0
1.0
10-15
-
-
15-20 60-200
Summary of the advantages and disadvantages of the various methods of blood sampling, Adept : Diehl, 2001 and NC3Rs. 2012
(
)
Mouse
Rat
Guinea
pig
Rabbit
Low
Low
4/Day
-
-
-
4/Day
4/Day
4/Day
-
Low
-
-
Repeat bleeds Time/Rest date
Route
Superficial temporal veins
Sapheneous/Pedal vein
Marginal ear vein
General
anesthesia
Tissue damage*
No
No
No(Local)
-
8/Day
2/Day
8/Day
Lateral tail vein
No
Low
No(Local)
8/Day
Central ear artery
Low
2/Day
8/Day
Jugular vein
Yes
Low
4/Day
4/Day
Tail tip amputation
Yes
Mod
No
No
No
Retrobulbar sinus/plexus
Yes
Mod/high
Cardiac, Axillary region,
No
No
No
No
Yes
Mod/high
Superior vena cava**
*The potential for tissue damage is base on the likely incidence of it occurring and the severity of any
sequelae, e.g. inflammatory reaction or histological damage.
**Only carried out as a terminal procedure under general anesthesia.
Anesthesia
Induce the state of unconsciousness
Reduce Pain and distress
Anesthesia
Inhalation anesthesia
• Diethyl ether
• Carbondioxide
• Isoflurane
• Halothane
Anesthesia
Injection anesthesia
• Barbiturates pentobabitone sodium “Nembutal”
• Ketamine: Xylazine
Injected Vol. (ml)= Dose (mg) x BW (kg) x ml
drug conc. (mg)
Euthanasia
“Is put to sleep animals”
Method of Euthanasia
I Physical methods
• Cervical dislocation
• Decapitation
• Shooting
• Electrocution
II Chemical methods
Anesthesia drug injection: Overdose
Gas or vapor Inhalation
• Toxic gas inhalation; CO2
• Volatile vapor inhalation
• Diethyl ether
Euthanasia
Cervical dislocation
Euthanasia
Cervical dislocation
Euthanasia
Decapitation:
Guillotine
Euthanasia
Diethyl ether and chamber
Euthanasia
CO2 Chamber
Euthanasia
Check Death
oNo heart-beat
oNo respiration action
oCheck for sign of muscular rigidity
oCheck papillary reflex
Necropsy Technique
o Check lesion in sick animal
o Collected organs for histology
o Health monitoring
Gastrointestinal Tract
Duodenum
Stomach
Colon
Jejunum
Cecum
ileum
Abdomenal Organ
Stomach
Liver
Spleen
Kidney
Reproductive organ
female
Seminal vescles
Ovary
male
Vas deferens
Prostate
Urinary bladder
Uterus
Testis
Reproductive organ
male
female
Thoracic Organ
Thymus
Lung
Heart
•
Diehl, K., Hull, R., Morton, D., Pfister, R., Rabemampiania, Y., Smith, D., Vidal, J., and
Vorstenbosch, C. 2001.A Good Practice Guide to the Administration of Substances and Removal
of Blood, Including Routes and Volumes. Journal of Applied Toxicology 21: pp. 15-23.
•
Joslin, J. 2009. Blood Collection Techniques in Exotic Small Mammals. Journal of Exotic Pet
Medicine. Vol 18. No 2: pp. 5117–139.
•
Lawson, P.T. 2000. Laboratory Animal Technician In AALAS Training Manual. Sheridan Book,
Inc., Chelsea.17, p.17-25.
•
NC3Rs. 2012.Blood sampling microsite. [Online]. Available:
http://www.nc3rs.org.uk/bloodsamplingmicrosite/page.asp?id=313.
•
Waynforth, H.B. and Flecknell, P.A. 1992. Experimental and Surgical Technique in the Rat.2nd
ed.Acardemic press. London and New York: pp. 69-89.
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