Animals
to be used in experimental protocols which involve extensive manipulation
should be handled frequently before the onset of the study to
allow the animal to acclimate to your scent making them more docile
while restrained. Within species, particular stocks or strains
of animal may have distinctive behavioral responses.
MICE
Adult
mice are picked up by the tail by compressing the base of the
tail between the thumb and forefinger and gently placing the animal
onto a solid surface. Rest the animal on the dorsal aspect of
your forearm and transport it short distances. Alternatively,
the animal can be placed in a small container with a cover containing
holes that admit in air. Adult mice can also be picked up by grasping
the loose skin over the shoulders and gently lifting the animal
from its cage or, alternatively, a pair of forceps (toothless)
can be used to grasp the mouse by either the tail or the skin
over the shoulders.
For
restraint the mouse is picked up by the tail as described above
and is placed over the wire bar lid of the cage and lowered until
the mouse grasps the wire with its forefeet. The excess skin over
the animal's back is grasped between your thumb and forefinger,
the hand is rotated so that the mouse is lying on its back within
the palm of the hand. The animal's head is closest to your thumb
while the tail is grasped with your smallest finger. The result
is a mouse that is immobilized for examination or manipulation.
Devices
are available to restrain mice for a variety of procedures. Commercially
available plexiglass restraining cylinders provide access to the
animal's tail for intravenous injection or blood collection. Home-made
devices can be made out of plastic syringe casings.
Conical
plastic sleeves, referred to as Decapicones®, can also be
used. The plastic is approximately the same thickness as that
of plastic waste bags. The flexible transparent clear-plastic
sleeve is conical, is open at its base, and has a small breathing
hole at the apex. The mouse is slid into the cone through the
base with its nose resting adjacent to the breathing hole. The
excess plastic is gathered and a rubber band is placed around
the animal's tail and the plastic of the cone. The cone permits
access to the tail and also, if the animal is positioned properly,
will permit parenteral injections through the thin-walled plastic.
Alternatively, the wire bar lid from a shoebox cage which contains
a food trough can be used to restrain a mouse to provide access
to its tail. The wire bar lid is set on a solid surface so that
it rests on the angular food trough. The mouse is directed between
the food trough and the end of the wire bar lid which is resting
on the top's surface. The mouse's tail is directed between the
wire bars and gently pulled so that the animal's rear end is held
firmly against the lid. This method provides access to the tail,
while limiting the mouse's ability to turn around and bite.
RATS
Rats
can be picked up by the base of the tail as has been described
for mice. However, extreme care must be exercised as an adult
rat's body weight is approximately 20 fold greater than an adult
mouse, whereas the tail is not 20 times greater in diameter. Therefore
it is much easier to injure the rat's tail. Common injuries include
fracturing coccygeal vertebrae (the small vertebrae within the
tail) or causing the skin to slip off the tail exposing underlying
tissue. It is essential that the rat be picked up by the base
of the tail as close to the body as possible. The rat should then
be placed on your forearm or a solid surface. A rat should not
be carried by their tail for more than a few seconds!
Alternatively
rats (< 300 grams) may be picked up by grasping the animal's
body from above so that the rats back is held firmly around the
thorax with your thumb and forefinger placed on either side of
the animals' head at the level of the mandible. When held firmly,
the rat is restrained and is unable to move its head to bite.
Large rats may be picked up similarly, however the hindquarters
must be supported with the other hand.
Like
for mice, there are larger commercial plexiglass rat restrainers
which provide access to the rats tail while protecting the handler
from the animal's head. Also larger Decapicones® are extremely
useful for handling rats. A larger cone is manufactured for this
species. They are utilized the same way as described for mice.
HAMSTERS
Males
are generally more docile than females. If simple precautions
are taken, hamsters can be routinely handled with minimal stress
to the animal and handler. Awakening the hamster from sleep will
frequently be met with an aggressive response. Hamsters can be
removed from their cage with the use of a small can or cup, which
they will usually enter; they can be scooped out with cupped hands;
or they can be grasped by the abundant loose skin over the dorsal
cervical region.
To
manually restrain a hamster begin by placing the animal down on
a solid surface. The palm of the hand is placed down over the
hamster with the thumb near the head. The excess skin is grasped
and gathered into your hand until the body wall is snug against
your fingers. The animal will be immobile and will not be able
to turn its head and bite.
GUINEA
PIGS
Guinea
pigs should be restrained using two hands. Your dominant hand
should be used to grasp the animal's thorax from below opposing
your thumb and fingers on either side of the animal's chest. The
second hand is used to support the hindquarters. For restraint
with greater control, the animal can be held using the same grip,
however the animal should be grasped from above the thorax instead
of below, and the pelvic limbs should be grasped and extended
while the animal is placed in dorsal recumbancy on a flat surface.
RABBITS
Appropriate
technique when handling rabbits is essential to prevent the animal
from accidentally traumatizing itself. Rabbits should be removed
from their cage by grasping the excess skin over the dorsal cervical
region. Rabbits should never be picked up by their ears. The rabbit's
hind end should be supported immediately after removal from its
cage. If not supported properly, the rabbit may kick with its
powerful hind limbs which may induce lumbar vertebral dislocation
or fracture (broken back). Rabbits can be moved for short distances
by permitting the animal to bury its head at the junction of your
body and your bent elbow, supporting the animal's body with your
forearm while putting gentle downward pressure over the animal's
back with your other hand.
Rabbit
restrainers are available that are extremely useful for transporting
or restraining rabbits. Rabbits are relatively easy to restrain
when using an appropriate restraining device. Rabbit restrainers
are usually of stainless steel, have a solid bottom, a top made
from wire bars, and contain a metal sliding plunger. Rabbits are
placed into the device by raising the wire bar top and pulling
the plunger away from the restrainer creating ample space into
which the rabbit is placed. The wire bar top is then closed and
locked into place, and the plunger slowly pushed in the direction
of the rabbit until it is gently but firmly displaced up the ramp
contained within the restrainer such that its nose is at the breathing
end of the ramp and the rabbit cannot move. This device provides
excellent access to the rabbit's ears, making it very useful for
blood collection or IV injections. The restrainer also provides
access to the animal's back for subcutaneous injections. Investigators
should sanitize the restrainer with appropriate
chemical agents before returning.
Restrainers
must be cleaned in a cage washer or suitably disinfected between
conventional and specific pathogen-free rabbits because of the
likelihood of spreading disease.
Rabbits
can also be restrained by using a towel. The towel is wrapped
around the animal's body such that it's head is covered and it
is swagged around the animals hindquarters. Either the ears or
the rabbit's dorsum is left exposed, dependent on the site that
requires access. Clean towels should always be used between different
groups of rabbits.
Back
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Parenteral
Injections
The
ability to administer materials by injection is essential for
most experimental studies employing laboratory animals. Anesthetics
and test compounds must frequently be administered to animal subjects
by injection. There are five commonly used routes of parenteral
administration: subcutaneous (SC), intraperitoneal (IP), intravenous
(IV), intradermal (ID), and intramuscular (IM). Not all techniques
are appropriate for each species. For example, IM injections are
avoided in the mouse because the amount of material that can be
injected into the mouse's limited muscle mass is so small that
the technique is not practical. IP injections are almost never
administered to rabbits, as other techniques are more suitable.
It
is essential that the appropriate parenteral site is selected.
Systemic absorption and distribution differ considerably between
sites. Dosage and volume of material administered must be carefully
considered relative to the type of agent, site of injection and
species used. The size of syringe and needle must also be considered.
In order to assure the delivery of an accurate volume of injected
material, the volume of the syringe should, in general, not exceed
the volume of material to be administered by 10 fold. The length
of the selected needle should be long enough that sufficient tissue
penetration is achieved but not be so long that it becomes unmanageable
or is likely to be inserted to far. The needle's size should be
as small (highest gauge) as possible to limit tissue trauma but
be large enough so that the injection can be made relatively rapidly
and without applying excessive pressure to the syringe plunger.
Syringe and needles should be of the locking type in order to
prevent accidental dislodgement which may result in autoinoculation
or back spray. Proper disposal of used needles and syringes is
essential. Needles should never be recapped, as the risk of accidental
injection is highest during recapping, and they should always
be disposed of into a designated sharps container.
Injection
volumes provided in this document are general recommendations.
Under some circumstances it may be inappropriate to inject the
recommended volume. For example, volumes should be reduced when
the agent is irritating or hypertonic. Volumes may be increased
when giving isotonic fluids for rehydration and fluid maintenance.
MICE
Subcutaneous
injection
SC
injections can be administered easily to mice. The needle is inserted
between the folds of skin into the base of the triangle that is
formed when traction is applied to the skin overlying the animal's
scruff. The syringe's plunger should be retracted to verify that
a vacuum is created and no blood or tissue fluid can be aspirated.
Subsequently, the plunger is depressed releasing the material.
In general no greater than 1 ml should be injected per SC injection
site in adult mice (> 25 grams). Several sites over the animal's
back should be used if larger volumes must be administered. In
general, needles should be 0.5 - 1Ó long and 23 ga or larger gauge.
Intraperitoneal
injection
The
administration of material into the peritoneal cavity is frequently
performed in mice. The aim of this technique is to administer
material into the space surrounding the abdominal organs, avoiding
injection directly into an organ. Mice should be restrained and
held with their ventrum exposed and head pointed downward, this
causes the freely moveable abdominal organs to move towards the
animal's diaphragm making accidental puncture of organs less likely.
A 1'' 23 ga or larger gauge needle is inserted into the abdominal
cavity in the lower right quadrant to avoid the cecum and urinary
bladder. The needle should be directed towards the animal's head
at an angle of 15 - 20 degrees and inserted approximately 5 mm.
Aspiration should be attempted to ensure that an abdominal viscus
(hollow organ such as the bladder or colon) has not been penetrated.
If material is aspirated, the syringe should be removed and disposed.
Never inject GI tract contents or urine into the peritoneal cavity,
as a bacterial or chemical peritonitis will likely result. In
general the volume of material administered into an adult mouse
should not exceed 1 - 2 ml.
Intravenous
injection
The
veins on the lateral aspect of the mouse's tail are an excellent
site for IV administration. The principal function of these veins
is for thermoregulation. They will dilate when the mouse's body
temperature rises in order to disseminate heat. Application of
heat to the whole animal or locally to the tail can be used to
cause venodilation making vascular access easier. The mouse should
be restrained so that its tail is accessible. A 0.5Ó 25 g or larger
gauge needle is used. The vein is located, the needle inserted
by directing the needle into the vein with its bevel pointing
upward at an angle of approximately 20 degrees. The needle is
inserted slowly visualizing the needle as it enters the vein.
Once the vein's wall has been penetrated the needle should be
directed cranially approximately 2 mm. Blood should be aspirated
into the needle's hub before making an injection.
During
material administration the vein should blanch and no material
or swelling should be detectable at the injection site. Material
should be administered slowly to avoid vascular overload or rupture
of the vein from excess pressure. No greater than 0.5 ml should
be administered IV to an adult mouse. Pressure should be applied
over the injection site by gently holding a cotton pledget or
piece of gauze over the injection site for approximately 30 seconds
to prevent hematoma formation. Preferably the needle should be
inserted into the vein midway down the tail, permitting additional
attempts for venipuncture proximally if the initial attempt is
unsuccessful.
RATS
Subcutaneous
SC
injections are performed in rats using the same technique as was
described for mice with the following differences. The volume
of material administered can be increased to approximately 5 ml
per site in an adult rat (>300 grams). Syringe size should
be increased proportionately and needles should be 22 ga or larger
gauge.
Intramuscular
IM
injections may be performed in the rat. Injection volumes are
limited to 0.25 ml site because of limited muscle mass. Either
the quadriceps muscles located on the cranial aspect of the femur
or the hamstrings on the caudal aspect of the femur can be used.
Care must be taken to avoid depositing material on or near the
sciatic nerve which runs along the caudal aspect of the femur
in the thigh. Therefore the needle should be directed cranially
if injecting the quadriceps or caudally when injecting into the
hamstrings. A 23 ga 0.5Ó needle or larger gauge should be used.
The needle is directed through the skin into the muscle belly
approximately 3-4 mm. Aspiration should be attempted before injecting
to determine that accidental penetration of a blood vessel has
not occured.
Intravenous
IV
injection technique for the rat is similar to the mouse. However,
the vessels are more difficult to visualize, especially in adult
rats. The skin overlying the vessels in adults becomes quite thick,
making vascular access much more difficult. For this reason the
preferred site for vascular access is near the tail base. Injection
volumes administered to an adult rat should not exceed 2 mls and
large volumes should be administered slowly to avoid vascular
overload. The technique describing IV administration and needle
size in mice should be followed.
Intraperitoneal
The
technique for IP injections in rats is virtually identical to
mice. Rats should be restrained with their abdomen exposed and
their head held downward. The injection site, method and needle
size is as described for mice. Because of their larger size <
5.0 mls of material can be administered to an adult rat.
HAMSTERS
Subcutaneous
SC
injections are performed in hamsters using the same technique
as was decribed for mice with the following differences. The hamster
has considerably more loose skin overlying the injecting site
permitting a proportionately larger volume of material to be administered
(< 3 ml). Syringe size should be increased proportionately
and needles should be 22 ga or larger gauge.
Intramuscular
IM
injections may be administered to the hamster. The technique is
as described for the rat except the injected volume is limited
to 0.15 mls site because of the hamster's limited muscle mass.
Either the quadriceps muscles located on the cranial aspect of
the femur or the hamstrings on the caudal aspect of the femur
can be used.
Intravenous
IV
injections are difficult to perform in the hamster because of
the lack of easily accessible veins. The OCV staff should be contacted
for additional information.
Intraperitoneal
The
technique for IP injections in hamsters is virtually identical
to described for mice and rats. Hamsters should be restrained
with their abdomen exposed and their head held downward. The injection
site, method and needle size is as described for mice. Because
of their larger size < 3.0 mls of material can be administered
to an adult hamster.
GUINEA
PIGS
Subcutaneous
SC
injections are performed in guinea pigs using the same technique
as was decribed for mice with the following differences. The volume
of material administered can be increased to approximately 5 mls
per site in an adult guinea pig. Syringe size should be increased
proportionately and needles should be 22 ga or larger gauge.
Intramuscular
IM
injections may be administered to the guinea pig. The sites and
methods are similar to those described for the rat. Injection
volumes can be increased to < 0.3 mls site because guinea pigs
have slightly larger muscles.
Intravenous
IV
injections are very difficult to perform in guinea pigs because
of the lack of easily accessible veins. The veins at the base
of the tongue or penile vessels (males) may be used. Consult the
OCV staff for additional information.
Intraperitoneal
The
technique for IP injections in guinea pigs is virtually identical
to mice. Guinea pigs should be restrained with their abdomen exposed
and their head held downward. The injection site, method and needle
size is as described for mice. Because of their larger size <
5.0 mls of material can be administered to an adult guinea pig.
Intradermal
injections
ID
injections may be used to immunize guinea pigs. In contrast to
SC injections where material is deposited into the space between
the skin and body wall, ID injections deposit material within
the layers of the skin. Therefore, the volume of material which
can be administered is very small (< 0.1 ml site; 0.05 ml recommended).
The fur should be clipped so that the injection site can be clearly
observed. A 0.5Ó 25 ga or larger gauge needle and a 1 cc syringe
are recommended. The area to be injected is swabbed with an alcohol
soaked cotton pledget, the needle is inserted bevel up into the
skin at approximately a 15 - 20 degrees angle. The needle is inserted
approximately 1 mm.
The
plunger is withdrawn to confirm there is significant negative
pressure and blood or fluid cannot be aspirated. The material
is injected slowly creating a small bleb that typically takes
several minutes to resolve. Immediate dissolution of the bleb
indicates that the material has been injected subcutaneously rather
than intradermally. ID injections should be made over the dorsal
thoracic and lumbar region. Multiple sites (up to 10) can be used.
RABBITS
Subcutaneous
injections
SC
injections are easily performed in rabbits because of the laxity
of their skin and the large area into which material can be administered.
The technique is the same as described for mice, however injections
should not be administered over the neck as this is the site from
which the animal is picked up. A 1Ó 22 ga or larger gauge needle
is recommended. Volumes should not exceed 5 mls per site unless
isotonic fluids are administered. Approximately 30 ml of isotonic
fluids can be administered per site and multiple sites may be
used. Always aspirate before injection.
Intramuscular
injections
The
recommended sites and technique for IM injection in the rabbit
are as described for rats. IM injection sites are illustrated
in. However, the rabbits larger muscle mass requires a longer
needle (1Ó) to adequately introduce the material deep within the
muscle belly, a distance of approximately 7 - 10 mm in an adult
rabbit and acceptable volumes to be injected are larger (<
1.5 mls).
Intravenous
injections
IV
injections are straightforward in rabbits because of the ease
of vascular access. The marginal ear veins located on the lateral
aspect of the rabbit's ears are readily visible. The veins can
be made more prominent by occluding the vessel at the base of
the ear by gently holding off with your fingers or applying a
small paperclip. The vein is swabbed with an alcohol soaked cotton
pledget. The needle is inserted as described for injecting rodent
tail veins. Remember to aspirate to verify placement of the needle
within the vein. Remove the paperclip or your occluding fingers
prior to injection. Volumes of approximately 5 to 10 mls can be
administered if given slowly, however routine volumes are frequently
< 1 ml. A butterfly needle or an Ïover the needleÓ catheter
may be inserted if larger volumes are to be administered or repetitive
injections will be given. A 0.5Ó 24 ga or larger gauge needle
is recommended. Pressure should be applied over the injection
site by gently holding a cotton pledget or piece of gauze over
the injection site for approximately a minute to prevent hematoma
formation. Intradermal injections
IntaDermal
Injections
ID
injections may be used to immunize rabbits. The technique is as
described for guinea pigs with the following changes. The neck
and anterior thoracic region should be avoided for injection as
rabbits are handled by grasping this region. Because of their
larger size up to 12 sites can be used.
Back
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Blood
Sampling
Collection
of blood from laboratory animals is frequently necessary for a
variety of experimental uses including determination of pharmacokinetics,
antibody production, clinical pathology evaluation, etc. Blood
may be collected from animals which are to survive the procedure
or at sacrifice as a terminal event. Whereas there is no limitation
on the amount of blood that may be collected terminally, the volume
collected from animals surviving the collection is limited to
prevent anemia and hypovolemia. As a general guide the 1-3-6 rule
should be followed. The rule states that the average blood volume
of most laboratory animals is 6% body weight (60 ml/kg); the most
blood that can be reasonably expected from a terminal sacrifice
is 3% body weight (30 ml/kg); and no more than 1% (10 ml/kg) body
weight may be collected during any 2 week period from animals
surviving the blood collection. Although venipuncture is generally
a satisfactory method for survival blood collection, catheterization
of a peripheral vessel may be necessary for animals requiring
frequent collection of small quantities of blood.
It
is extremely important to apply pressure to the blood collection
site, especially when penetrating an artery, for several minutes
post blood collection to prevent hematoma formation.
MICE
There
are a variety of methods that are utilized to collect blood from
mice. The techniques described below are recommended by OCV staff
for survival (tail vein or orbital venous sinus) or terminal blood
collection (cardiac).
Lateral
tail vein (Saphenous v.) venipuncture
The
veins located on the lateral aspect of the mouse's tail are useful
for collecting small volumes (< 0.1 ml) of blood. The technique
for venipuncture is as described for IV injection except that
a small volume of blood is aspirated into the syringe instead
of injecting material. The use of a needle without a syringe,
allowing the hub to fill with blood, and subsequent collection
into a microhematocrit tube is useful when very small quantities
of blood are needed.
Orbital
venous sinus collection
The
sinus surrounding the globe of the mouse's eye is a useful site
for collecting larger volumes of blood from surviving animals.
The schematic provided in illustrates the location of the sinus.
General anesthesia must be provided when collecting from this
site. Under general anesthesia the mouse is grasped so that its
back rests on the palm of your left hand (right hand if you are
left-handed) with its head toward your thumb. The thumb is placed
just just lateral to the animal's trachea so that the jugular
vein on the same side as the eye from which you are collecting
blood is occluded and the fur on the animals head is drawn into
the palm of your hand. This causes the animals eye to proptose
(bulge) slightly. Be careful not to occlude the trachea! A 50
ul microhematocrit tube which has been broken in two is directed
into the medial canthus (junction of eyelids closest to the animal's
nose) of the eye rotating slightly as the tube is directed to
a point directly behind the globe, inserting the non-broken end
first. Sufficient pressure must be applied to cut through the
fibrous layer which surrounds the sinus. Blood flows through the
tube and occasionally around the tube once the sinus has been
penetrated. After blood collection, the tube is removed and the
eyelids closed and a dry cotton pledget is applied over the eye
with gentle pressure to prevent retroorbital hemorrhage. In general
blood should not be collected from the same eye more than 3 times,
allowing at least 1 week between collections. An antibiotic opthalmic
ointment may be applied following bleeding.
Cardiac
puncture (diaphragmatic approach)
Cardiac
puncture is the preferred technique for terminal collection of
large blood volumes. General anesthesia is administered and the
animal placed on a solid surface with its ventrum exposed The
xyphoid process is palpated at the caudal aspect of the animal's
sternum. A notch is present on both sides of this process. A 1Ó
22 ga or larger gauge needle attached to a 1 - 3 ml syringe is
inserted into either notch and directed toward the heart as determined
by palpating for the apex beat. Negative pressure should be applied,
by placing slight backward pull on the plunger, once it has been
inserted beneath the skin. Reflux of blood is apparent once the
needle has penetrated the heart. The animal must be sacrificed
at the completion of the procedure prior to awakening from anesthesia.
RATS
Lateral
tail vein (Saphaneous v.) venipuncture
The
procedure for collecting blood from the rat's tail vein is similar
to the technique described for the mouse. A slightly smaller gauge
needle (24 ga or larger gauge) can be utilized. Because of the
vein's size, larger blood volumes (approximately 1 ml) may be
obtained from adult rats.
Orbital
venous plexus
The
technique describing blood collection from the mouse's orbital
venous sinus should be followed for orbital venous plexus collection
in the rat. The only difference in that the vessels surrounding
the rat's globe are a network of small veins rather than a blood
filled sinus and the fibrous connective tissue surrounding the
plexus is quite dense. Therefore, the broken end of the hematocrit
tube which serves as a cutting edge should be inserted into the
plexus. Remember this technique must be performed under general
anesthesia and post bleeding hemostasis is essential to prevent
complications.
Cardiac
puncture (diaphragmatic approach)
The
technique for cardiac puncture from the rat is identical to that
described for the mouse except a longer (1.5Ó) and smaller gauge
(22 ga) needle is recommended. A 5 - 10 cc syringe should be used
if large blood volumes are desired. This procedure is performed
as a terminal event only and general anesthesia is required. The
animal must be sacrificed at the completion of the procedure prior
to awakening from anesthesia.
HAMSTERS
Orbital
venous sinus
The
technique describing blood collection from the mouse's orbital
venous sinus should be followed for orbital venous sinus collection
in the hamster except the microhematocrit tube should be inserted
into the lateral canthus rather than the medial. The technique
must be performed under general anesthesia and post bleeding hemostasis
is essential to prevent complications.
Cardiac
puncture (diaphragmatic approach)
The
technique for cardiac puncture from the hamster is identical to
that described for the mouse except a smaller gauge (22 ga) needle
is recommended. A 3 - 5 cc syringe should be used if large blood
volumes are desired. This procedure is performed under general
anesthesia as a terminal event only. The animal must be sacrificed
at the completion of the procedure prior to awakening from anesthesia.
GUINEA
PIGS
Orbital
venous plexus
The
technique describing blood collection from the rat's orbital venous
plexus should be followed for orbital venous plexus collection
in the guinea pig. Remember this technique must be performed under
general anesthesia and post bleeding hemostasis is essential to
prevent complications.
Cardiac
puncture (diaphragmatic approach)
The
technique for cardiac puncture from the guinea pig is identical
to that described for rats. This procedure is performed as a terminal
event only and general anesthesia is required. The animal must
be sacrificed at the completion of the procedure prior to awakening
from anesthesia.
RABBITS
Central
auricular artery
The
central auricular artery is a useful site for collection of moderate
volumes of blood from rabbits which are to survive the procedure.
Vasodilatation should be induced by administering the phenothiazine
tranquilizer and alpha adrenergic receptor blocker acepromazine
(0.25 - 0.5 cc SC) approximately 5 - 10 minutes prior to blood
collection. A 21 ga or larger gauge butterfly needle is preferred,
however a 1Ó 21 ga or larger gauge needle and syringe may also
be utilized. The insertion site is disinfected using an alcohol
soaked pledget prior to inserting the needle, bevel up, into the
artery. An immediate flashback is observed and blood is allowed
to flow out of the open end of the butterfly needle into a suitable
container or alternatively blood can be collected directly into
a syringe. It is essential to apply pressure to the artery over
the insertion site for at least 3 minutes to provide suitable
hemostasis. Significant blood loss can occur from the artery if
adequate hemostasis is not provided.
Cardiac
puncture (diaphragmatic approach)
Large
volumes of blood can be collected directly from the heart of anesthetized
rabbits as a terminal event. The technique is similar to that
described for mice and rats, however a larger needle and syringe
(1.5Ó >18 gauge ; >20 cc) should be used. Death must be
confirmed at the completion of the procedure by administering
pentobarbital (120 mg/kg) IV.
Table
I
Recommended
Needle Sizes and Injection Volumes for Various Parenteral Techniques
Route
|
Gerbil
|
Hamster
|
Mouse
|
Rat
|
IM
|
Quadriceps,gluteals
0.1 per site
<23 gauge |
Quadriceps,gluteals
0.1 per site
<23 gauge |
Quadriceps
0.03 per site
<23 gauge |
Quadriceps,gluteals,triceps
0.2-0.3 per site
<22- gauge |
IP
|
Lower
right quadrant
of abdomen
2-3
<21 gauge |
Lower
right quadrant
of abdomen
3-4
<21 gauge |
Lower
right quadrant
of abdomen
1-3
<21 gauge |
Lower
right quadrant
of abdomen
10
<22 gauge |
IV
|
Lateral
tail veins
0.2-0.3
<23-gauge |
Not
Recommended |
Lateral
tail veins
0.2-0.3
<23-gauge |
Lateral
tail or saphenous veins
0.5 - 3 slowly
<22-gauge |
Intra-
gastric
|
Stomach
NA
18-22-gauge,3-4 cm long
Bulbed feeding needle |
Stomach
NA
18-22-gauge,4-4.5 cm long
Bulbed feeding needle |
Stomach
5-10 mL/kg
18-22-gauge,2-3 cm long
Bulbed feeding needle |
Stomach
5-10 mL/kg
15-18-gauge,6-8 cm long
Bulbed feeding needle or
8 french flexible catheter |
SC
or SQ |
Neck,back
2-3
<21 gauge |
Neck,back
3-5
<21 gauge |
Neck,back
2-3
<22 gauge |
Neck,back,abdomen
5-10
<21 gauge |
|
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