Name
______________________________
LABORATORY ANIMAL MANAGEMENT
ANATOMY AND CLINICAL TECHNIQUES
Contents:
I.
II.
III.
IV.
Directions
Directions
Directions
Directions
for
for
for
for
Lab
Lab
Lab
Lab
1
2
3
4
V.
Handling and Restraint
VI. Ear Notching
VII. Injections
VIII. Blood Collection
IX. Oral dosing/stomach tubing/gavage
X.
Scalpel Use
XI. Suturing
XII. Anesthesia
XIII. Euthanasia
XIV. Dissection
XV. Clean up
XVI. Ear Notch Code
XVII. Anatomy—structures to identify
XVIII.Anatomical terms for direction
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Bring directions for labs
1 – 4 to class. You should
also bring a copy of the
anatomy structures to ID.
The remaining sections are
to further explain the
directions. You may print
them or not as you wish.
The information is
connected by hyperlink.
Use control click to move
from the highlighted terms
to more detailed
explanations.
1
I.
Lab 1:
Safety, animal handling, needle use, suturing.
A.
Station 1: Rats (Some will be sedated by instructor)
1.
Practice tickling: This is a method of gentling
the rat. You play with the rat as they would play
with each other which gets them used to being
handled
and
makes
them
less
afraid
of
manipulations used in the lab.
2.
Practice sexing
3.
Practice handling and restraint techniques (in
hands, against body, in pocket).
B.
Station 2: Mice (Some will be sedated by instructor)
1.
Practice sexing.
2.
Practice handling and restraint techniques (in
hand and on cage top).
C.
Station 3: Needle use
1.
Practice putting together the needle and syringe
and filling the syringe to the appropriate level.
2.
Practice injection techniques using an orange.
3.
Practice SQ and IP injection restraint and
placement using a stuffed animal and a dry needle
and syringe.
4.
Use Koken rat to practice tail vein injections
(use the 28 g needle with attached syringe, inject
with dI water only).
D.
Station 4: Scalpel use
1.
Practice putting the blade
taking the blade off safely.
on
the
handle
and
E.
Station 5: Suturing
1.
Practice opening and closing hemostats.
Ideally,
hemostats should be held by your thumb and 3rd
finger with the 2nd finger wrapped around the
finger loop and the 1st finger providing support to
the shaft.
2.
Practice suturing on a glove-wrapped foam pad.
Please make sure someone checks your technique.
F.
Station 6:
Anatomy—Study pictures and rabbit model.
 ALL DISPOSABLE NEEDLES, SYRINGES, AND SCALPEL BLADES MUST GO
INTO THE SHARPS CONTAINER. SAVE SUTURE NEEDLES.
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II.
Lab 2:
Anatomy and Clinical techniques (It is recommended
that you bring a full page copy of the anatomy terms from the
back of this manual on which to take notes.
I have also
provided a PowerPoint Dissection Guide from which to study.)
A.
Each group will be provided with 1 each euthanized rat
and mouse. We will go over all procedures together.
B.
Injections
1.
Practice restraint techniques for these procedures
and get comfortable handling the equipment using
both the rat and the mouse.
a)
SQ
b)
IP
c)
Ear notch
C.
Dissection: Follow along with the instructor. Do not
work ahead or you may miss techniques or structures.
1.
Note external characteristics of the male and
female rat and mouse.
2.
Make a mid-line incision from neck to genitalia.
Use the scalpel blade for the rat and scissors for
the mouse. Separate skin from underlying muscle.
3.
Observe the structures in the neck.
Practice
collecting blood from the jugular vein by
inserting a needle through the pectoral muscle.
4.
Make a mid-line incision through the peritoneum of
the abdominal cavity careful not to cut through
the xiphoid process. Cut laterally at the cranial
and caudal ends of your incision to open the
peritoneal cavity.
5.
Observe the in situ position of the abdominal
organs. The instructor will go over each system.
6.
Enter the thoracic cavity by cutting through the
ribs on either side of the sternum and removing
the sternum.
A wider opening may be made by
cracking the ribcage.
7.
Observe the in situ position of the thoracic
organs and go over them with the instructor.
8.
The instructor will demonstrate the method of
decapitation using bone shears on the rat and
scissors on the mouse. We will then dissect and
identify the structures of the brain and head.
 WASH INSTRUMENTS IN NOLVASAN USING A BRUSH. RINSE THEM, DRY
THEM AND PUT THEM AWAY AS DIRECTED
 ALL NEEDLES AND SYRINGES, SCALPEL BLADES, AND MICROHEMATOCRIT
TUBES MUST GO INTO THE SHARPS CONTAINER
 ALL CARCASSES, ANIMAL TISSUE, AND ITEMS WITH BLOOD ON THEM,
MUST BE BAGGED FOR INCINERATION
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III. Lab 3--Clinical techniques
A.
Rats:
Each group obtains 1 rat and practice the
following:
1.
Gavage: Attach a gavage needle to a dry syringe.
Measure the needle against the size of the rat.
Dip the tip of the needle into sucrose solution.
Restrain the rat in your non-dominant hand against
your body.
Insert the gavage needle into the
rat’s mouth and down the esophagus up to the hub
of the needle, and then remove it slowly.
2.
SQ: Restrain the rat against your upper body to
deliver 0.2 cc of saline subcutaneously.
3.
IP: Deliver 0.2 cc of saline.
4.
Ear notch: Restrain the rat against your upper
body and practice ear notching.
C.
Follow directions in the manual for anesthesia.
1.
Calculate the appropriate dosage and administer
the anesthesia.
2.
Fill out log sheet to monitor anesthesia.
3.
Apply eye lubricant to each eye, using a piece of
clean gauze, to prevent drying.
4.
Inject 10 cc/kg (1 cc/100 g bwt) of sterile saline
SQ
to
prevent
dehydration
caused
by
the
anesthesia.
You will not need to use the
restraint technique on the anesthetized animal.
5.
Warm a microwavable heating pad (heated 1 - 1½
minutes, 2 minutes for 2 pads) and cover it with a
towel.
Place the anesthetized animal on the pad
to
maintain
body
temperature
while
under
anesthesia and to facilitate vasodilatation for
blood collections.
D.
Site preparation:
Shave the rat ventrally, dorsallaterally, on the dorsal surface of the hind legs, and
on the proximal surface of the tail.
E.
ID: Using a 27 g needle, inject 0.05 cc – 0.1 cc/site
of saline between the layers of the skin on the dorsallateral surface of the rat. Done correctly, a bleb, or
blanching bubble, should form.
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F.
Lateral saphenous vein:
1.
Use a rubber band and hemostat to create a
tourniquet at the upper leg joint.
2.
Use a 23 g needle held perpendicular to the leg to
puncture through the skin and into the vein.
Remove the needle.
Collect 100 µl of blood from
the surface of the skin using a microhematocrit
tube.
G.
IV—Tail vessels (Primary focus—rat vein):
1.
To collect from the tail vein, use a rubber band
and hemostat to create a tourniquet at the base of
the tail. Insert a 25 g needle into the vein and
collect 100 µl of blood from the hub of the needle
into a microhematocrit tube.
2.
You may also try collecting from the tail artery,
which lies laterally from the anus to the tip of
the tail. Don’t use a tourniquet on the artery.
H.
Cardiac puncture: Use the xiphoid process as your
landmark for point of entry. Use a 23 g, 1-inch needle.
Hold the needle at a 20 to 30° angle and direct
smoothly into the heart. Collect 0.1 cc to demonstrate
correct placement and technique.
I.
Euthanize the rat using CO2
1.
Study the anatomy—be sure you can identify all of
the structures on the list at the end of this
packet.
Look at other animals to observe
anatomical variation and the structures of both
sexes.
2.
Practice suturing on the rat skin and abdominal
muscle.
IMPORTANT:
 Wash instruments in Nolvasan using a toothbrush.
dry them and put them away as directed.
 ALL NEEDLES AND SYRINGES, SCALPEL BLADES, AND
SUTURE NEEDLES MUST GO INTO THE SHARPS CONTAINER.
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Rinse them,
DISPOSABLE
5
IV.
Lab 4: Comparative Handling and Clinical Techniques
A.
Station 1:
Mice: Techniques are similar to the rat;
however, the mouse may be restrained in one hand.
1.
Gavage: Use an appropriate sized gavage needle.
2.
SQ:
Restrain the mouse against a cage top to
deliver 0.1 cc of saline.
3.
IP: Deliver 0.1 cc of saline.
4.
Ear notch: Scruff the mouse and ear notch.
5.
Lateral saphenous vein: Restrain the mouse in an
adapted syringe case. Pinch the leg in the groin
to extend the leg and to occlude blood flow. Use
small clippers to shave the outside of the leg.
Use a 23 g needle held perpendicular to the leg to
puncture through the skin and into the vein, then
remove the needle and collect the blood pooling on
the surface.
Collect 100 µl of blood in a
microhematocrit tube.
6.
Submandibular vein: Use a 5 mm lancet to puncture
the blood vessels located at the junction of the
mandible and maxillary cheek bones.
Collect 100
µl of blood from the surface of the cheek into a
microhematocrit tube.
7.
Cardiac
puncture:
This
technique
requires
anesthesia; alternatively, blood may be collected
immediately after euthanasia.
Use a 25 g, 5/8”
needle.
Use the xiphoid process as your landmark
for point of entry. Hold the needle at a 20 to 30°
angle and direct smoothly into the heart.
B.
Station 2: Rabbit
1.
Handling and sexing
a)
Practice picking up the rabbit, holding the
rabbit in a “football” hold, and returning
the rabbit to its container.
b)
Practice flipping the rabbit.
Identify the
sex of the rabbit and note the external
genitalia (on both sexes if available).
2.
SQ Injection:
Inject 1 cc of sterile saline
anywhere along the dorsal side (back) of the
animal using a 20 g needle and a 1 cc syringe.
3.
Blood collection:
Collect 100 µl of blood from
the marginal ear vein (auricular vein).
C.
Station 3: Guinea pigs and/or hamsters
1.
If these species are available, practice handling
and sexing these animals.
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V.
Handling
A.
Mice and rats—handling
Back to Lab 1
1.
Pick animals up by the base of the tail. That is
the side of the tail closest to the body. Picking
them up by the tip of the tail makes them feel
insecure, they are more likely to bite, and there
is the potential for stripping off the skin of the
tail if animals are handled this way.
B.
Mice—restraint
Back to Lab 1
1.
Mice are scruffed in the non-dominant hand.
2.
Hold the mouse by the tail using your dominant
hand.
This is usually done on the cage top; it
should always be done on a rough surface.
3.
Using your non-dominant hand, slide your thumb and
first finger (or knuckle) on either side of the
mouse’s neck and down along the jaw line.
Close
your fingers grasping the skin, and then pull it
taut up over the back of the head and neck of the
mouse.
You should have enough skin so that the
head cannot turn.
4.
Position the body of the
mouse against your thumb.
Use your second finger to
pull skin from the mouse’s
back
taut
against
the
thumb.
5.
Grasp the tail between
your little finger and the heel of your hand. The
mouse should be supported all along its body. The
tail should always be restrained. If the tail is
loose, it will pull the mouse’s body loose.
C.
Rat—restraint
Back to Lab 1
1.
Rats are restrained using the
“V” technique.
Place your
first and second finger on
either side of the rat’s head
and wrap your thumb and last
2
fingers
under
the
forelimbs.
The hand used
depends on your goal; learn
the technique in both hands.
Always
support
the
hindquarters
when
using
this
technique. Adult rats are too
heavy to hold in one hand.
You may use your other hand or hold the rat
against your body for this additional support.
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2.
3.
4.
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To restrain for a subcutaneous injection, you will
position the rat perpendicular to your body
against your chest.
a)
Grab the rat in a V hold in your non-dominant
hand.
b)
Pull the rat away from your body, grab the
tail, wrap the tail to the top of your nondominant arm, and place the rat against your
chest so that it is parallel to the floor.
Press the rat against your body along its
length using your hand and arm.
c)
Place your thumb on the rat’s shoulder blade.
Move your index finger over the rat’s head to
the far side of its body, and pinch the skin
up between your fingers to make a tent.
Remember to continue pushing down on the
rat’s head with your fingers as you pinch.
To restrain the rat for gavage, use the V hold in
your non-dominant hand.
a)
Stretch the rat out by grasping the tail and
hindquarters and pulling them away from your
body so that the rat’s hind feet are not
underneath it, and then positioning the rat
vertically against your chest. Use your hand
and arm to press the rat into your body.
b)
In this hold, your first two fingers will
pull gently on the jaw line of the rat
elevating its head and straightening out its
body. The thumb and last 2 fingers may pull
the forelimbs back as well.
The goal is to
achieve a straight body so that the feeding
tube may slide down the esophagus unimpeded.
To restrain the rat for an intraperitoneal
injection, use the pocket technique.
Start with
the rat in the V hold in your dominant hand.
a)
Position
the
rat
head down and tail
up over your nondominant
pocket.
Pull your pocket out
with
your
nondominant
hand
and
slide the rat down
into the pocket.
b)
Transfer your hold from your dominant to nondominant hand by placing your non-dominant
hand over the top of the pocket and pressing
the rat into your hip or leg.
Slide your
8
c)
D.
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dominant hand out of the pocket as your nondominant hand rolls up to cover the rat.
Slide the non-dominant hand up to the rump of
the rat so that you may use your thumb to
push the rat down into the pocket. Continue
to push the rat into your body and cup your
fingers around the rat’s body so that it
can’t turn in your pocket.
Position the leg and tail between your thumb
and hand and roll the rat’s hindquarters away
from your body to visualize the abdomen.
Rabbit—handling
Back to Lab 3
1.
Pick the rabbit up by the scruff; however, this is
not as cranial a position as it is on the mice.
It is easiest to grasp the skin over the shoulder
blades.
You may use either hand, but it is most
comfortable in your dominant hand. When you have
a firm grip on the scruff, place your other (nondominant) hand under the hindquarters and lift the
rabbit. Once the rabbit is out of
its container and in your arms,
wrap your non-dominant hand around
the rabbit’s body, placing the
rump in your hand and tucking the
rabbit’s head under your arm. This
is called the “football” hold and
is used for transporting rabbits
for short distances.
2.
When returning rabbits to their cage or box,
always put them in rump first and head towards
your body.
This is to prevent them from jumping
out of your arms, which could result injury.
3.
Rabbits may be flipped to visualize their ventral
side.
a)
Scruff the rabbit with your dominant hand and
pull the rabbit away from your body.
b)
Using your non-dominant hand, roll the rump
around your dominant arm, then bring the
rabbit back to your body so that the belly is
up and the rabbit is caught between your
dominant arm and your body.
c)
You may hold the rabbit on your dominant arm
to examine the head and teeth. Use your hold
on the scruff to pull the head back.
d)
You may transfer the hold to the non-dominant
arm to examine the abdomen and hind feet and
to sex the rabbit.
Wrap your non-dominant
9
arm around the rabbit and place its rump into
your hand.
To sex the rabbit, pull down on
the tail.
E.
F.
G.
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Rabbit restraint
1.
Rabbits do not generally require a firm restraint
for subcutaneous injections. We place the rabbit
in a box to keep in a small space.
2.
Rabbits are placed into a restrainer for more
invasive procedures such as blood collections.
This is important to
prevent
the
rabbit
from trying to jump,
and
in
doing
so,
breaking
its
back.
We will use a cat bag
to
restrain
the
rabbit.
Hamster handling
Back to Lab 3
1.
Hamsters have almost no tail and so all handling
and restraint must be done using a full body
scruff.
2.
Press the hamster into
the cage with your nondominant hand.
Grasp
the skin along the back
between your thumb on
one
side
and
your
fingers on the other.
Make sure you have the
skin along the back and
behind the head pulled
taut so that the hamster cannot turn within its
skin. Then lift the hamster.
Guinea pig handling
1.
Guinea pigs are never
scruffed.
They
are
picked up by wrapping one
hand around their body
about the level of the
shoulder blades. Use the
other hand to scoop up
the hind quarters and
lift the guinea pig.
10
VI.
Ear Notching
Back to Lab 2
A.
Use the chart in back to learn the Ear Notch Code for
Cole B. This is not a universal numbering system, but
we will use this system for both rats and mice in this
class. Note that the chart is looking from the back of
the animal’s head with the right ear on the right.
B.
Mice
1.
Restrain your mouse using the scruffing technique
described above (use your non-dominant hand).
With the head immobilized and the tail restrained,
place the mouse onto a cage top with the mouse
pressed up against the right side of the cage top.
If done correctly, the mouse should be unable to
turn its head to the right.
2.
Grasp the ear notcher in your dominant hand
holding near the tip of the ear notcher.
3.
Slide the tip of the ear notcher over the margin
of the mouse’s ear.
Depress the tip of the
notcher to clip a ½-circle of tissue from the
margin of the mouse’s ear. (For some numbers, you
will have to move toward the center of the ear and
clip a whole circle.)
C.
Rats
1.
Restrain the rat in a V-hold in your non-dominant
hand.
Hold the rat against your body, pressing
the head close to your body. You may wrap the rat
in a towel for this restraint if needed.
2.
As with the mice, slide the tip of the ear notcher
over the margin of the ear and clip a ½-circle of
tissue from the ear.
VII. Injections
Back to Lab 1
A.
Needle safety website:
http://safetyservices.ucdavis.edu/snfn/safetynets/snml/
sn3/SN3pdf
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B.
Needle use
Needle Size Recommendations for Class Procedures
Needles
20 g, 1-inch
23 g, 1-inch
Rat
SQ
IP
IC
L. Saphenous
Mouse
L. Saphenous.
Rabbit
SQ
25 g, 5/8-inch
IV (tail blood
collection)
SQ
IP
IV (tail)
IC
IV (ear vein)
27 g, 1/2-inch
28 g, 1/2-inch
1.
2.
3.
4.
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ID
IV (tail vein injection) IV (tail vein injection)
-
Capping needles:
To cap a needle lay the cap on
the table and slide the needle into the cap. If a
firm seal is desired, lift the syringe and press
into the table to seal the cap.
Always lay
needles in the cap when setting them on the table—
it keeps the needles clean and prevents accidents.
Beveled edge:
Insert the needle with the angled
surface up to direct the needle parallel to
vessel.
Aspirate: Draw back on plunger to check location
of needle.
a)
For anything other than an IV procedure,
aspiration of blood is an indication of an
improperly placed needle.
You may have
entered an organ, in which case injection of
fluid may cause tissue damage.
If blood
appears, the needle should be pulled out and
repositioned.
Note:
Aspiration is not
necessary for an ID injection.
b)
If urine or feces are aspirated (yellow,
green, or brown), the needle, syringe and
solution
are
contaminated
and
must
be
discarded to prevent infection.
Disposal: Needles and syringes must be discarded
into a Sharps container. Do not cap needles prior
to disposal.
Do not remove the needle from the
syringe.
Have a Sharps container close to where
you are working.
12
C.
Syringe use
1.
Milliliter (ml) and cubic centimeter (cc) are
equivalent
volumes
and
may
be
used
interchangeably.
2.
For
injection,
chose
a
syringe
that
will
accurately measure the volume you need.
You can
only measure to one 100th of a ml in the 1 or 0.5
cc syringe and even then not very accurately.
Dilute solutions for more precise measurements.
3.
For blood collections, use small syringes for
small blood vessels or choose a collection device
that uses capillary action (microhematocrit tube).
Too much vacuum will collapse the vessel.
4.
Read the syringe where the black edge of the
plunger touches the liquid in the syringe.
5.
When inserting the syringe into the animal, keep
your thumb off of the plunger to avoid premature
delivery.
Move your thumb to the plunger after
aspirating.
D.
Subcutaneous (SQ):
Used in all rodents and rabbits.
Used primarily for slow release dosing as with fluid
replacement or hormone administration. Rabbits are
often given sedatives and anesthetics subcutaneously.
Ease of administration makes up for the slightly slower
uptake.
Back to Lab 3
1.
General technique
a)
Usually given in scruff of neck, but when
multiple sites are needed, injections may be
given along the back.
b)
Skin should be tented, or pinched up away
from the body, and the needle positioned
close to the body but between the skin and
the underlying muscle.
c)
Aspirate.
Resistance
indicates
correct
positioning.
If the plunger pulls back
easily, you have probably gone through the
skin and out the other side.
2.
Mice are scruffed (always hold the tail), and then
placed in a ventral position on a cage top. Use a
25 g, 5/8-inch needle.
Deliver 0.1 cc of saline
to practice.
3.
Rats are held horizontally against the body using
the hand, wrist, and arm of the non-dominant hand.
Use a 23 g, 1-inch. Deliver 0.2 cc of saline to
practice, or give 10 cc/kg to hydrate during
anesthesia.
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4.
Rabbits are placed in a box; they do not require
restrain for this procedure. Pinch the skin to
make a tent anywhere over the back. Use a 20 g,
1-inch needle. Inject 1 cc of saline.
E.
Intraperitoneal (IP): Used in all rodents; can be used
in the rabbit, but the restraint is difficult. Uptake
is more rapid than SQ.
Back to Lab 3
1.
General technique
a)
Hold the animal with its head down.
This
allows the abdominal organs to fall cranially
decreasing the risk of incorrect penetration.
b)
This injection is given in the lower abdomen
about the level of the knee joint on the
right side of the animal’s body (this may be
to your left).
c)
This injection is going into the abdominal,
or intraperitoneal, cavity.
Position the
needle at about a 30 degree angle from the
surface of the body. Do not insert parallel
to the body or your needle may penetrate the
skin, but not the muscle layer.
d)
Aspiration is particularly important.
(1) Blood indicates improper placement into
an organ. Remove needle and reposition.
(2) Urine
or
feces
indicates
improper
placement into the bladder or GI tract.
Remove needle and place needle and
syringe into the sharps container.
2.
Mice are scruffed and held head down. Use a 25 g,
5/8-inch needle. Deliver 0.1 cc of saline to
practice the injection.
3.
Rats are restrained using the pocket technique.
Use a 23 g, 1-inch needle.
Deliver 0.2 cc of
saline to practice or the appropriate dose of
anesthesia.
F.
Intradermal (ID):
Used in antibody research, the
injected material is usually an antigen, or foreign
protein, coupled with an adjuvant, a material used to
increase
irritation,
thus
producing
an
antibody
response. Tuberculin tests use this type of injection.
ID injections are performed primarily in GP and
rabbits, but we will practice on an anesthetized rat.
1.
Hair is shaved off of the back, lateral to midline
to avoid injecting over the spinal cord.
2.
Use a 27 g, ½-inch needle and a 1 cc syringe.
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14
3.
4.
5.
The skin is pinched between the thumb and finger
forming a tent. Hold the syringe
at the hub and brace the index
finger of each hand together to
allow for a strong force with a
small,
controlled
movement.
Slide the needle along the top of
the tent inserting it within the
layers of skin. Once the needle
is inserted, drop the skin so that
you can visualize the delivery.
Deliver 0.05 – 0.1 cc of sterile
saline per injection site.
Try
several sites on the back.
A small bubble that blanches white
will appear at the injection site.
This bubble is called a bleb.
Back to Lab 4
G.
Intramuscular (IM):
Seldom done in the mouse or rat,
common site in guinea pigs and rabbits. Muscle masses
commonly used are quadriceps muscle of the leg or
lumbar muscles of the back. We will not practice this.
H.
Intravenous (IV): Often difficult on rodents, but very
common in rabbits.
Used for very rapid absorption,
usually medicinal. Practice on the Koken rat. You may
also practice on your anesthetized mouse and rat if you
wish.
Back to Lab 1
1.
Always inject into veins.
An injection into an
artery may damage tissue as the fluid injected
passes through the small capillaries at the distal
end of the appendage. An injection into a vein
travels through the large vessels to the heart and
is diluted before reaching the capillaries.
2.
Use a 28 g needle attached to a ½ cc syringe and
inject 0.1 cc of saline into the tail vein of the
rat and mouse.
3.
Hold the tail in your non-dominant hand across the
first three fingers and under your little finger
to form a plateau. Squeeze the tail gently between
your thumb and first two fingers to stabilize.
4.
Always direct the needle towards the body.
5.
Hold the syringe at the hub and with fingers on
top and sides, not underneath as this raises the
syringe and puts the needle in at too steep an
angle.
Use only the fingers to slide the needle
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15
6.
7.
in.
Stabilize your hand, by placing the little
finger against the fingers holding the tail.
Do not aspirate. Inject the saline and look for
clearance of blood along the vein. A bubble
forming in the tail at the site indicates that you
are not in the vein.
Make your first injection about half way down the
tail and move up the tail as you make further
attempts (distal to proximal).
VIII. Blood Collection:
Back to Lab 4
A.
General procedures
1.
Vasodilatation
a)
This facilitates blood flow reducing stress
to both the animal and the technician. It may
be accomplished with chemical (acepromazine)
or
mechanical
means
(heat
or
topical
irritants).
b)
We will produce vasodilatation with heat.
(1) Obtain a microwavable gel pack and heat
in the microwave (1½ minutes)
(2) Place the animal on the pack, with a
towel between the pack and the animal. A
second towel may be used to cover the
animal.
(3) The temperature within the toweling
should be about 30ºC. Rapid breathing,
panting or drooling are indications of
hyperthermia.
If this occurs, remove
the rat from the heating pad.
2.
Site preparation
a)
For most blood collections, you will clip the
hair prior to beginning.
We will use
clippers, but you may also use a scalpel
blade or a depilatory.
It is not necessary
to clip prior to cardiac puncture.
b)
If hair is clipped, site is usually cleansed
to remove debris that may be picked up in the
sample. However, excessive cleaning will
cause irritation so limit it to a quick wipe
with dry gauze or gauze wet with EtOH or warm
water or use masking tape to pick up hair.
3.
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When collection is complete
a)
Remove the needle, and the tourniquet, if one
is used.
b)
Press a clean gauze pad over the point of
entry. Apply firm, but gentle pressure. Do
16
c)
d)
B.
Blood collection guidelines from IACUC campus policies:
http://safetyservices.ucdavis.edu/ps/a/IACUC/po/bloodVo
lumes
1.
2.
C.
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not hold so tight as to stop blood flow to
the site.
Hold continuously for 30 seconds. Do not dab
or wipe during this time.
Observe the animal for an additional 30
seconds to assure that bleeding has stopped.
1% of body weight collected every 2 weeks. (Body
weight is in grams; blood volume in milliliters.)
Exp: 200 g rat x 1% = 2 ml
Lateral Saphenous vein:
Anesthesia is not required,
but may be used.
BACK
1.
Mice may be restrained in a
20 cc syringe case; rats may
be restrained in a towel.
2.
Occlude the blood vessel.
a)
Mouse: The hind leg is
extended
and
held
between the thumb and
Mouse
finger.
Pinching
the
skin in the groin helps
to extend the leg and to
occlude the vein.
The
hair
is
shaved
using
clippers.
b)
Rat:
Wrap the rubber
band once around the leg
and catch the ends of the
band with the hemostats. Rat
Twist
the
hemostats
to
tighten
the
band
and
occlude the vein.
3.
The lateral saphenous vein runs
over the top of the foot, along
the outside of the calf muscle
just above the ankle, and up the back of the leg.
4.
A 23 g, 1-inch needle is inserted perpendicularly
into the muscle to puncture the
saphenous vein.
The needle is
removed and blood is collected
from the surface of the skin in a
100 l capillary tube. Collect no
more than 100 l to demonstrate
17
5.
success. Then cover the puncture with a gauze pad
and compressed until bleeding stops. Flexing the
foot may also help.
If the first puncture is not
successful, move closer to the
foot and try again.
Blood flows
from the foot up the leg, so if
the
vein
is
damaged,
moving
upstream
is
more
likely
to
produce a successful blood collection.
D.
Submandibular area—Mice only: Blood flows better in an
unanesthetized mouse, but the technique requires a very
secure restraint.
You may choose to try it with or
without anesthesia. BACK
1.
Directions with pictures:
2.
http://www.medipoint.com/html/mouse_phlebotomy.htm
l
3.
http://www.youtube.com/watch?v=niTVnEAHOko
4.
Scruff the mouse and orient it so that you are
looking at the cheek. Locate the junction of the
mandible and maxillary cheek bones. This is also
the junction of the lower facial vein and the
submandibular vein.
The actual location is not
easy to see so we will use the “freckle” as our
landmark and move slightly dorsal and caudal from
that mark.
5.
Using a 5 mm lancet, puncture the skin at that
point. Although not visible through the skin, the
concentration of vessels makes it relatively easy
to find and penetrate a vein.
6.
Blood
will
pool
on
the
cheek.
Use
a
microhematocrit tube to collect blood.
7.
Cover the puncture with a gauze pad and compress
until bleeding stops.
E.
Tail vessels: Anesthesia is not required, but we will
use to prevent pain to the animal while you are
learning a difficult procedure.
This technique is a
relatively non-invasive way to collect blood from the
mouse or rat, but it takes practice, particularly in a
pigmented animal.
BACK
1.
Vasodilatation--the animal must be warm.
If you
have not already done so, place your rat on a
towel-wrapped microwavable gel pack for 5-15
minutes until the veins in the tail are distended.
Continue to work with rat on the hot pack during
the blood collection.
18
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2.
3.
4.
5.
6.
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Positioning and tourniquet:
a)
To collect from one of the tail veins, place
the rat in lateral recumbency (on its side).
Make a tourniquet by wrapping a rubber band
once around the tail, grasping both ends with
a hemostat, and twisting the hemostat several
turns.
This allows the tourniquet to be
easily removed.
b)
To collect from the tail artery, place the
rat in dorsal recumbency (on its back) and
look for the artery on the ventral surface of
the tail from the anus to the tip of the
tail. Do not use a tourniquet.
Using your non-dominant hand,
drape the tail across your first
three fingers and under your
little finger to form a plateau.
Squeeze the tail at each side
gently between your thumb and
first two fingers to stabilize.
Use a 25 g needle without a
syringe.
Holding the needle by
the sides of the hub, insert the
needle beveled edge up.
It
should be parallel to the tail
and very superficial.
If blood
appears in the hub of the needle, use a
microhematocrit tube to collect a maximum of 100
µl.
If you fail to get blood, move
a short distance to try again.
a)
When collecting from the
vein, move distal (away
from
the
body)
because
blood flow comes from the
tip of the tail.
b)
When
collecting
from
the
artery,
move
proximal (toward the body) because blood flow
is coming from the body.
Once you fill your microhematocrit tube, remove
the needle, and the tourniquet, and use a gauze
pad to apply pressure until bleeding stops.
19
F.
Cardiac Puncture:
Always done under anesthesia and a
terminal procedure on mice, rats, and rabbits, it
allows for collections of large quantities of blood
(approximately 5 to 6% of body weight or 5 - 6 ml/100 g
bwt in the rat). Use a 23 g, 1 to 1 ½-inch needle and
a 1 cc syringe for the rat, a 25 g, 5/8-inch needle for
the mouse.
Back to Lab_4
1.
Palpate the notch between the xiphoid process and
the last rib. This is your point of entry.
2.
Insert the needle into the notch,
parallel to midline and at a 30
angle.
Insert smoothly; hesitation
may cause the needle to deflect the
heart.
Insert until the hub is
slightly underneath the last rib
3.
Pull back on the plunger to form a vacuum.
If
blood does not flow into the syringe, keeping the
plunger pulled out about 0.1 cc, pull the needle
out slowly. If the needle went through the heart,
it may reenter.
If blood
still doesn’t flow, remove
the needle completely and
start again. Keep the needle
moving in a straight line.
Excess movement in the chest
cavity could lacerate organs.
4.
Collect a minimum of 0.1 cc
to demonstrate correct placement.
Then, remove
the needle.
5.
This is a terminal procedure due to the risk of
thoracic bleeding.
G.
Auricular (Ear) vessels (rabbit only)
BACK
1.
Blood collections do not require sedation, but a
sedative will make the procedure easier on both
the animal and the technician.
Acepromazine is
commonly used because it provides vasodilatation
as well as sedation.
A topical analgesic is
often used, either in place of, or in addition to
the sedative.
2.
The marginal ear vein can provide up to 5 cc. We
will practice this technique, collection 100 µl
per attempt.
a)
Clip hair to improve visibility.
b)
Apply prilocaine, a topical analgesic. Allow
this to sit on the ear about 5 minutes. Then
wipe it off before inserting the needle.
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20
c)
3.
IX.
Use a paper clip on gauze placed proximal to
the point of entry as a tourniquet.
d)
Hold a roll of gauze under the ear to
stabilize.
e)
Insert a 25 g needle into the vein.
Needle
should be inserted parallel to the ear; the
vein sits above the surface of the skin.
f)
Pull blood from the hub of the needle into a
microhematocrit tube for small volumes.
g)
If larger volumes are needed, you would use a
22 g needle and a small syringe.
Draw back
slowly to prevent the vein from collapsing.
The central artery can be used for volumes up to
50 ccs. We will not attempt this technique in
class.
a)
Don't use a tourniquet.
b)
Use a 20 g needle with a Vacutainer tube or
break the hub off of the needle and allow
blood to drip from the end of the needle into
a test tube.
Oral dosing/stomach tubing/gavage
Back to Lab 3
A.
This is a technique used to deliver fluids directly
into the stomach. It allows for a measured dose to be
delivered at a specified time. It is also used when
solutions are unpalatable or degraded by enzymes in the
mouth.
B.
A stomach tube (feeding tube or gavage needle) is a
needle with a ball on the end. A properly sized needle
will have a ball which allows easy passage into the
esophagus but which is too large to pass through the
larynx into the trachea.
C.
Feeding tubes come in straight or curved and in
stainless steel or plastic.
We will use plastic
because it tends less traumatic to the animal, but it
is possible for the animal to chew the tube.
Try to
prevent this by not hesitating once you have inserted
the tube.
However, this is not always possible to
prevent. If chewing occurs, the tube may be rough and
cause trauma to the esophagus or the tube may break and
the animal may swallow part of it. If the tube becomes
rough, replace it. If the animal swallows part of the
tube, let me know. It may regurgitate the tube, but if
not we will euthanize the animal.
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D.
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Technique for stomach tubing--You may practice this
technique on a lightly sedated animal but not on an
animal that has been heavily sedated.
The muscles of
the trachea in a heavily sedated or anesthetized animal
are too relaxed to prevent entry of the tube into the
lungs, which could result in trauma.
1.
Attach a gavage needle to an empty syringe.
2.
Measure the tube.
The needle should extend from
approximately the tip of the nose to the last rib.
3.
Dip the tip of the needle into the sucrose
solution to trigger the swallowing reflex and to
increase the animal’s acceptance of the tube.
4.
Restrain the rat and hold against your body
leaving your dominant hand free.
The mouse does
not need to be held against your body—a single
hand hold is sufficient.
5.
Insert the tube into the
mouth by sliding it into
the
diastema
and
then
pulling it to the front of
the
mouth.
Straighten
your
first
2
fingers
against the animal’s jaw
to pull its head back.
Use the tube as a lever to
tip the animal's head back
forming a straight line
from the mouth to the
esophagus.
6.
Pass the tube along the
palate
to
the
larynx.
Time
entry
into
the
esophagus
with
the
swallowing reflex.
7.
Slide the tube down the
esophagus
into
the
stomach. When the hub of the needle is at the
mouth, the end should be in the stomach.
8.
If you were delivering a sample, you would depress
the plunger on the syringe at this point to
deliver the correct dose.
9.
Remove the needle carefully.
Observe the animal
for signs of distress.
Resistance on the needle or struggling from the animal
could mean improper placement. Remove the tube and try
again.
22
X.
Scalpel Use
Back to Lab 1
A.
Loading and unloading the scalpel blade onto the
handle.
1.
Open the package and pick up the blade with a pair
of hemostats. Position the blade in the hemostats
so that the blade faces away from
your hand.
2.
Slide the opening of the blade over
the raised portion of the handle
until it clicks into position.
3.
To unload, turn the handle over so
the
raised
portion
faces
the
counter. Hold the handle over the
sharps container. Use the hemostats
to pop the blade off into the sharps container.
B.
Scalpel grip
1.
Pencil grip--hold the scalpel handle as a pencil.
This allows for greater precision over short
distances. Most appropriate grip for our work.
2.
Fingertip grip--hold the scalpel handle between
fingertips and thump tip.
Allows for greater
control with a longer range of motion.
May be
more comfortable for rat dissections.
C.
Incision technique
1.
Use one hand to exert tension on the skin
longitudinally and laterally
to
stretch
the
cutting
surface by placing the first
finger and thumb on either
side of the incision line.
This hand will move as the
blade progresses to keep
enough tension to see the
cutting edge.
2.
The scalpel blade is place behind (not between)
the fingers. The blade slides in a straight line.
Enough pressure should be exerted to cut through
the skin, but not the underlying muscle layer.
Try not to lift the scalpel blade until the
incision is finished.
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XI.
Suturing
Back to Lab 1
A.
Suture needles
1.
Round needles are used for soft tissue.
2.
Cutting needles with a triangular cross-section
are used for tough tissues like skin.
B.
Suture material
1.
Non-absorbable suture is used primarily for skin
where it can be removed. If left in the body, it
will become encapsulated in fibrous tissue.
The
most common material is braided silk although
nylon and even stainless steel may be used.
2.
Absorbable suture loses tensile strength and then
breaks down and is absorbed by the body, usually
within 60 days. Examples include catgut, chromic
gut, and Vicryl.
These are used for internal
sutures.
C.
Suture size
1.
Largest diameter 5 (0.7 mm) to smallest 11/0 (0.01
mm) (11/0 is pronounced 11 “ought”)
2.
In mice and rats is 3/0 or 4/0 is most common.
D.
Hemostat
1.
The
loops
of
the
hemostat
should be around your thumb and
3rd finger.
2.
Your first finger guides the
tip. Your second finger guides
the base.
E.
Simple interrupted sutures
1.
Position the needle in your left-hand hemostat.
Hold the needle between the center of the curve
and the swag (threaded end). Pick up one side of
the incision with a pair of rat-toothed forceps
and insert the needle in, taking a “bite” of about
5-8 mm. Pick up the opposing side of the incision
directly opposite the first and insert the needle
in it.
(You can sometimes pick up both sides at
once but watch for slipping.)
2.
Switch to a hemostat in your right hand, use it to
grasp the needle and pull it through the tissue
holding your left hand hemostat against the
incision as a brace if needed.
Pull the suture
material until only 2-3 cm is left on the left
side of the incision.
3.
Let go of the needle and move the hemostat in your
right hand to a point in the suture material about
10 cm from the right side of the incision.
Alternatively, you may use your hand to hold the
suture material instead of a hemostat.
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24
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
F.
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Wrap the suture material in your right hand (long
end) around the closed tip of the hemostat in you
left hand twice forming two loops.
This wrap is
counter-clockwise; over the top and to the center.
Grab the short end of the suture material with the
hemostat in your left hand.
Pull the short end of suture through the two loops
and tighten by crossing your right hand over your
left; continue in the same direction as your wrap.
Do not let go of the suture in your right hemostat
(or hand) or change the position of your hands at
this point.
Let go of the suture in the left hemostat and
staying “inside the V,” wrap the long end of
suture around the hemostat once.
This wrap is
clockwise; over the top and toward the center.
Using the left hemostats, again grasp the loose,
or short, end of suture. Pull it through the loop
and uncross your hands to tighten the knot (again
continuing in the same direction as your wrap).
You have now completed a square knot
Do not let go of the suture in your right hemostat
(or hand) or change the position of your hands.
To make a locking stitch, let go of the suture in
the left hemostat and staying “inside the V,” wrap
the long end of suture around the hemostat once.
This wrap is counter-clockwise.
Pick up the short end of suture, pull it through
the loop and cross your hands to tighten.
Cut the suture ends to about 5 mm in length. Make
your next stitch about 8-10 mm away.
You may reverse this procedure, by driving the needle
in from right to left, however, you will have to wrap
the opposite directions and cross left over right to
tighten the knot.
1.
Always wrap over to under.
2.
Always tighten in the same direction in which you
are wrapping.
25
XII. Anesthesia
A.
Drugs we will use for pain management
Drugs
Acepromazine
Ketamine
Lidocaine 1%
Prilocaine
Xylazine
B.
Actions
Sedative, Vasodilator
Dissociative anesthetic
Local anesthetic
Topical analgesic
Sedative, Analgesic
Muscle relaxant
BACK to Lab 4
Commercial Concentration
10 mg/ml
100 mg/ml
10 mg/ml
25 mg/g
20 mg/ml
100 mg/ml
Drug dosages
1.
Drug dosages are given in a formulary put out by
various veterinary organizations. Dosages are
usually expressed as a range and may vary from one
formulary to another. Your choice within the
range may depend on the procedure, as well as the
line, age, sex, etc., of the animal.
2.
Formulary dosages are expressed in mg/kg. The
actual delivery dosage will be in ml/kg and must
be calculated based on the commercial
concentration of the drug (from above table). The
drugs are sold in differing concentrations so read
the label carefully to be sure you have the
correct concentration.
Formulary dose x 1/conc. = delivery dose
Species/
Drug
Formulary dose Inverted
Delivery
Use
Used in lab
Commercial
Dose
Conc
Rat & mouse/ Acepromazine 0.5 mg/kg
1 ml/10 mg
0.05 ml/kg
Sedation
Rat & mouse/
Clinical
Ketamine
Xylazine
(100 mg/ml)
Acepromazine
Rat/Surgical
3.
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Ketamine
Xylazine
50 mg/kg
1 ml/100 mg
0.5 ml/kg
5 mg/kg
1 ml/100 mg
0.05 ml/kg
0.5 mg/kg
1 ml/10 mg
0.05 ml/kg
1 ml/100 mg
1 ml/20 mg
0.9 ml/kg
0.45 ml/kg
90 mg/kg
9 mg/kg
Note that some of the delivery doses are too small
to accurately measure when calculated for an
animal that weighs less than 1 kg.
26
C.
Dilution
1.
Drugs often come in concentrations too high to
accurately measure when drawing up volumes for
small rodents. We correct for this by diluting
drugs with sterile water, saline, or another
appropriate vehicle. The drugs we use in this
class will be diluted with sterile water.
2.
Acepromazine is diluted to different volumes for
rats and mice because of their 10-fold weight
difference.
Concentration of drug x volume concentrated drug/total volume = diluted concentration
Diluted concentration (inverted) x formulary dose = diluted delivery dose
Species/
Use
Rat
sedation
Mouse
sedation
Commercial
Conc.
10 mg/ml
Dilution
factor*
1:10
Diluted
conc
1 mg/ml
Formulary
dose
0.5 mg/kg
Del Dose
(dilute)
0.5 ml/kg
10 mg/ml
1:100
0.1 mg/ml
0.5 mg/kg
5 ml/kg
*Dilution factor = 1:n+1 where n+1 equals total volume
Dilution ratio = 1:n (ratio of solute to solvent)
D.
Drug combinations for balanced anesthesia
1.
Acepromazine alone (above):
Instructors will use
in lab 1 to produce light sedation for handling of
mice or rats. This drug will calm the animal but
the animal will remain conscious.
2.
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Ketamine:Xylazine:Acepromazine: This cocktail will
provide
anesthesia
for
clinical
techniques.
Ketamine
is
an
anesthetic.
Xylazine
and
acepromazine are sedatives. Adding sedatives will
calm the animal thereby reducing the amount of
anesthesia required.
Ace is also a vasodilator
making it easier to collect blood. In the table
below, we are mixing drugs and diluting with water
to make a cocktail that will provide balanced
anesthesia
in
an
easily
measurable
dosage.
Animals will lose consciousness and will not feel
pain; however, they may not lose the toe pinch
reflex.
27
Set delivery dose
Target Dilution:
Dilution Factor:
Cocktail volume:
dilution factor.
so all drugs are equal in volume
divide delivery dose by formulary dose
Multiply target dilution by concentration
Determine the required total volume and divide by the
Drug
Delivery
dose
Ket
Xyl
Ace
Water
1 ml/kg
50 mg/kg 1 ml/50 mg 100 mg/ml
1 ml/kg
5 mg/kg
1 ml/5 mg
20 mg/ml
1 ml/kg
0.5 mg/kg 1 ml/0.5 mg 10 mg/ml
Water or solvent needed to reach total volume
3.
Target
dilution
Concentration
Dilution
Factor
2
4
20
Total
volume
20 ml
10 ml
5 ml
1 ml
4 ml
Calculate the delivery dose of the pre-mixed
cocktail for rats.
a)
Weight the rat and convert to kilograms
Kg = g bwt/1000
b)
4.
Formulary
dose
Multiply the weight in kg times the delivery
dose of 1 ml/kg
1 ml/kg x __________ kg = ________ ml
Mice:
Mice require a higher dilution of the
ket:xyl:ace cocktail since a typical mouse weighs
about 30 grams and the rat dose would be
1 ml/kg x 0.03 kg (30 gram) = 0.03 ml which is
impossible
to
measure
accurately
with
our
syringes. We will dilute the rat anesthesia 1:10
for mouse anesthesia.
The dose will be 10 ml/kg
(0.1 ml/10 grams) so the delivery dose for a 30 g
mouse will be 0.3 ml.
a)
Weigh the mouse in grams.
b)
Calculate the delivery dose of the pre-mixed
cocktail for mice:
0.1 ml/10 grams x _______ grams = ________ ml
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5.
Ketamine plus xylazine (rats): This will provide
a surgical plane of anesthesia for surgical
procedures in lab 6.
Ketamine is again the
anesthetic.
We will continue to use xylazine as
the sedative and a muscle relaxant to prevent
trauma caused by surgical manipulations.
Both
will be at higher dosages to maintain a prolonged
period
of
anesthesia.
We
will
not
use
acepromazine as vasodilatation is contra-indicted
in a surgical procedure.
In this case, we are adding 1 ml of xylazine to a 10 ml bottle of ketamine.
Drug
Delivery
dose
Formulary
dose
Target
dilution
Concentration
Dilution
Factor
Ket
Xyl
1 ml/kg
1 ml/kg
90 mg/kg
9 mg/kg
1 ml/90 mg
1 ml/9 mg
100 mg/ml
100 mg/ml
1.1
11
Total
volume
11 ml
10
1
E.
Draw up dose and administer the anesthetic:
1.
Use a 1 ml syringe with the appropriate needle—23
gauge for rats, 25 gauge for mice.
2.
Note:
If your calculations give you a final
volume of more than 1 cc, please check your
calculations.
3.
Once you have completed your calculations, see the
instructor or TA for the anesthesia.
4.
Inject the anesthesia IP.
5.
Record dose and time on the anesthesia log sheet
provided in lab.
F.
Monitoring anesthesia: Monitor the depth of anesthesia
throughout the procedure and administer a supplemental
dose if the animal becomes light.
1.
Pedal withdrawal or toe pinch reflex:
The best
method of monitoring rodents for a surgical plane
of anesthesia is by using the spinal cord reflex
of the toe.
When you pinch the toe (using your
fingernail or a pair of forceps), the foot
withdraws away from the stimulus. Since reflexes
are reflexes are suppressed from head to toe under
anesthesia, the toe pinch reflex is one of the
last reflexes lost.
Note:
It is not necessary
for this reflex to be suppressed for clinical
techniques and it may not disappear with the light
anesthesia.
It should disappear with the heavy
anesthesia used for surgery in lab 6.
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29
2.
3.
4.
5.
Purposeful movement or vocalizations, those in
response to touch or painful stimuli, indicate
that the animal is light.
Muscle tone: An increase in tone, as demonstrated
by chewing or whisker movement, means animal is
light.
Breathing patterns:
Rapid and shallow if light;
slow and gasping if deep.
Color of mucus membranes (O2 levels)--blue or clear
if too deep.
G.
Supplemental doses:
Although the loss of the pedal
withdrawal reflex is our major sign that the animal has
reached a surgical plane, it may not disappear under
the light anesthesia dose. The animal should be
unconscious and unresponsive to stimulus before you
begin your procedures.
If your rat does not achieve
this level from the initial dose, or if it becomes
light during your procedures, supplement with ketamine
only (no acepromazine or xylazine unless directed to by
the instructor or TA).
1.
Supplemental
Ketamine:
This
should
be
administered at a dose of 1/3 to 1/2 of the
original dose.
Use a 1/3 dose when toe pinch
reflex is slight.
Use a 1/2 dose when toe pinch
reflex is strong or animal still exhibits the
righting reflex.
You may need to use the 1/2 cc
syringe to accurately measure this dose for
smaller animals. Syringe units: 50 units = 0.5 ml
Low dose (1/3 x 0.9 ml):
0.3 ml/kg
High dose (1/2 x 0.9 ml): 0.5 ml/kg
H.
Record keeping:
Record all drugs administered on the
Anesthesia Log (provided in class).
5.
Surgical records are required by the AWA and PHS.
2.
Ketamine is a controlled substance (Schedule III)
and records of its use are required by the FDA.
XIII. Euthanasia
A.
Ask for assistance in using the CO2 chamber.
If you
prefer, you may ask the instructor or TA to euthanize
your animal for you.
If you do not need to use the
euthanized animal, leave it in a cage with a cage top
on the back counter and we will euthanize the animals
in a group.
B.
Place the animal to be euthanized in the CO2 chamber
located in the back of the room.
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C.
D.
E.
Turn on the gas using the large knob on top of the tank
to start the flow.
Turn counterclockwise.
Then use
the small round knob on the regulator to release the
gas into the chamber.
Euthanasia will take 2-3 minutes for a mouse, 3-5
minutes for a rat.
Look for cessation of breathing.
Leave the animal in
the chamber for 1 minute after all signs of life have
disappeared.
If the animal is not being dissected,
insert a scalpel blade into the thoracic cavity
(thoracic punch) to assure death.
Back to Lab 4
XIV. Dissection:
BACK to Lab 2
A.
Use the following websites to prepare for lab and to
study for tests. There are also several good books on
anatomy at the Health Sciences Library.
See the
recommended resources on the class website for some
suggestions.
1.
Virtual necropsy:
http://tvmouse.compmed.ucdavis.edu/
2.
Labeled dissection:
http://www.utm.edu/staff/rirwin/public_html/RatAna
t.htm
B.
Locate the organs and landmarks in the abdominal and
thoracic cavities and in the neck that are found on
your list of anatomical structures to identify.
Note: The heart will beat for several minutes
following death. This is because the sinoatrial node
will stimulate beating as long as there is ATP in the
cells.
XV.
C.
Obtain the bone shears to decapitate your rat or ask
for help with this procedure.
Examine the brain and
find the structures on your list.
D.
Look at animals from other groups to note anatomical
variation between sexes and individuals.
Clean up.
BACK
A.
Dispose
of animal carcasses and tissue in the
appropriate bag for incineration.
Include any bloodcontaminated material, i.e., gloves or gauze, in the
bag.
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B.
Dispose of sharps (disposable suture needles, injection
needles and syringes, scalpel blades) in Sharps
container.
C.
Scrub your instruments using Nolvasan and a brush.
Rinse and dry them. Then sort them as directed.
DO NOT PUT WET INSTRUMENTS AWAY. They will rust.
D.
If a heating pad has been used, wash it with either
hand soap or Nolvasan, dry it, and return it to the
Styrofoam container.
E.
Return your lab coat and any
appropriate bag to be laundered.
F.
Wash your hands before leaving.
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cloth
towels
to
the
32
XVI
Back to
Ear Notching
Ear Notch Code for Cole B
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XVII Anatomy: Be sure that you can identify the following structures in dissected animals, the
rabbit model, and in diagrams.
Back to Lab 2
Blood vessels
Reproductive--female
Thoracic cavity
Lateral saphenous vein
Cervix
Lungs
Tail vein (mouse/rat)
Uterine horns
Heart
Tail artery (mouse/rat)
Oviduct
Diaphragm
Auricular artery (rabbit)
Ovary
Thymus
Reproductive-male
Neck
Peritoneal cavity
Preputial gland
Salivary glands
Xiphoid Process
Testes
Trachea
Stomach (Pyloric)
Epididymus
Esophagus
Stomach (Cardiac)
Gubernaculum
Thyroid
Duodenum
Pampiniform plexus
Masseter muscle
Pancreas
Vas deferens
Small Intestine
Seminal vesicles
Head
Cecum
Coagulating gland
Cerebral hemispheres
Large intestine
Prostate gland
Cerebellum
Liver
Penis
Pineal gland
Auricular vein (rabbit)
Gallbladder (mouse/rabbit)
Olfactory nerves
Urinary Bladder
Optic nerve & chiasm
Kidney
Hypothalamus
Adrenal gland
Pituitary (Ant. & Post.)
Spleen
Harderian gland
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XVIII
ANATOMICAL TERMS FOR DIRECTION
Cranial
Caudal
Toward the head
Toward the tail
Dorsal
Ventral
Toward the back or top
Toward the abdomen
Back to Lab 2
Anterior Toward the head
Posterior Opposite to the head
Superior
Inferior
Above
Below
Medial
Lateral
Toward the middle or midline of the body
Toward the side or away from the middle or midline
Proximal
Distal
Nearer the long axis of the body or a reference point
Away from the long axis of the body or reference point
Palmar
Plantar
Relating to the palm of the forelimb
Relating to the sole of the hindlimb
Oral
Concerning the mouth
Rostral
Toward the nose
Recumbency: Position in which an animal is lying
Sternal recumbency: lying on its abdomen
Lateral recumbency: lying on its side
Dorsal recumbency: lying on its back
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Superior
Dorsal
Cranial
Anterior
Caudal
Posterior
Superior
Anterior
Cranial
Inferior
Ventral
Ventral
lateral
medial
Proximal
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Dorsal
Distal
Caudal
Posterior
36
Inferior
Crossword puzzle to study anatomical directions—Answers are posted
separately.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
Across
Down
1.
3.
6.
8.
11.
12.
14.
15.
16.
2.
4.
5.
7.
9.
10.
13.
Toward the tail
Relating to the sole of the hindlimb
Relating to the palm of the forelimb
Away from the midline of the body
Concerning the mouth
Closer to a specific point on the body
Above
Toward the head
Toward the rear
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Further from a specific point on the body
Toward the abdomen
Toward the midline of the body
Toward the front
Toward the nose
Toward the back
Below
From Techtalk
Vol.15, No.4
Aug 2011
37