- Oral or per os (po)- - through the mouth
- Gavage - - into the stomach via a feeding
needle
When drugs, vaccines, injectable anesthetics or other agents are to be
administered, one of several different routes may be selected. The route
is governed by the nature of the agent being administered, the animal, and
the purpose of the administration, among other factors.
Oral: Substances
may be administered orally by addition to the food or the drinking water.
The amount of drug consumed by an animal will vary between each
individual. If the drug imparts an unpleasant taste, affecting
palatability, the drug will be consumed in decreased amounts.
Environmental conditions such as the ambient temperature will also affect
both water and food consumption. The general rule of thumb is that 5 grams
of food and 10 mls of water will be consumed daily per 100 grams of body
weight.
Gavage: If it is
necessary to administer exact amounts of a substance gastric feeding
needles should be used. Entry normally may be obtained without anesthesia
using hand restraint. Feeding needles with a ball tip helps prevent
introduction of the needle into the trachea and prevents trauma to the
oral cavity (Figure 1).
Flexible or plastic tubes may be bitten or chewed and are not recommended
for rodents. Feeding needles are inserted through the mouth into the
stomach or lower esophagus. Placing the needle next to the rat so that the
end of the needle is adjacent to the last rib and marking the position on
the needle that is adjacent to the tip of the nose will determine the
distance the feeding needle should be advanced into the oral cavity to
insure administration of the compound into the stomach (Figure
2). Care must be taken that the tube or needle does not enter the
trachea or puncture the esophagus or stomach, therefore knowledge of the
oropharyngeal anatomy is necessary. In most cases, introduction of the
tube toward the rear of the mouth will induce swallowing and the tube will
readily enter the esophagus. A violent reaction (coughing, gasping)
usually follows accidental introduction of the tube into the larynx or
trachea. With the mouse restrained in one hand the feeding needle is
introduced in the space between the incisors and the beginning of the
molars (diastema)(future picture). If introduced from the left diastema
the needle should be directed caudally toward the right ramus of the
mandible. As the needle approaches the pharynx the mouse will usually
swallow allowing introduction into the esophagus. Using the feeding needle
to gently extend the neck facilitates introduction into the stomach
(future video). With the stomach tube fitted to a syringe or aspirator,
materials may be administered or withdrawn as required.
Parenteral
Administration
- Intravenous (iv) - - directly into the vascular system through a
vein
- Intra-arterial (ia) - - directly into the vascular system through an
artery
- Intraperitoneal (ip) - - into the abdominal cavity
- Subcutaneous (sc) - - under the skin
- Intramuscular (im) - - into a muscle
- Intradermal (id) - - between layers of skin
Parenteral routes of
administration involve injections into various compartments of the body.
Sites used for collection of blood from veins may also be used for
intravenous administration. Intraperitoneal administration is one of the
most frequently-used parenteral routes in rodents. Other locations are the
musculature and subcutis. Materials given intramuscularly must be in very
small volumes and is generally not recommended unless necessary.
Absorption by this route is more rapid than from subcutaneous
administration. Regardless of the route used, it is essential that the
subject be securely restrained to prevent unnecessary struggling by the
animal and to avoid injury to personnel by dislodged needles.
The investigator should know the physiological and chemical properties
of the substance that he/she plans to inject. Considerable tissue damage
and discomfort can be caused by irritating vehicles, drugs or solutions
when injected into animals. The use of the foot pad as an injection site
for antigens with or without adjuvant is discouraged since it is a
needless and painful procedure. More suitable sites for antigen injections
are subcutaneously in the axilla or lateral thoracic wall, deep in large
muscle masses, or into the popliteal lymph node.
Equipment: 20-25 g needle, 1-3 ml syringe, rat holder,
warming lamp.
Volume: The volume injected IV into an adult rat
should not exceed 0.5 ml.
The lateral veins (future picture) of the tail are the
most frequently-used veins. Best results are obtained if the tail is
immersed in warm water for 5 to 10 seconds or the rat warmed for 5 to 15
minutes in the cage with a warming lamp with a 40 to 100 watt bulb. The
veins can be seen when the tip of the tail is lifted and rotated slightly
in either direction. The tip of the needle can be followed visually as it
penetrates the vein. Trial injection verifies proper needle placement.
Also, accurate placement can be confirmed when the vessel is visually
flushed when the compound is administered. The formation of a bleb at the
site indicates improper placement of the needle. A second attempt can be
performed by removing the needle and trying a site on the same vessel in a
more proximal location on the tail. Practice is essential. Younger rats
are easier to inject since the skin tends to thicken in adults making
accurate needle placement difficult.
The lateral saphenous vein on the lateral aspect of
the hindleg may also be used for intravenous injection. Light anesthesia
and/or confinement in a cylindrical rat holder is usually necessary. Other
sites that have been utilized are the sublingual vein and dorsal penile
vein in anesthetized rats.
Prolonged IV administration/sampling can be accomplished by jugular or
femoral vein catheterization, requiring surgical implantation.
Equipment: Syringe and 20-25 g, 1 to 1.5-inch needle,
preferably with a short bevel.
Volume: The volume injected IP into an adult rat
should not exceed 10 ml.
The rat is grasped as
previously described,
and held in dorsal recumbency in a head-down position. The injection is
made in the lateral aspect of the lower left quadrant (Figure 4).
The use of a short bevel needle inserted through the skin and musculature
and immediately lifted against the abdominal wall, aids in avoiding
puncture of the abdominal viscera. Utilizing the needle cover which has
been cut 1 to 1.5 cm from the end will limit the depth of injection.
Immobilizing the left leg is also essential in reducing this risk.
Restraint is best accomplished with a second person holding the rat in a
head-down stretched-out position; light anesthesia is recommended. Rapid
injection, especially with a large syringe, may cause discomfort and
tissue damage and should be avoided.
Equipment: Syringe and 20-25 g, 1/2 to 3/4-inch
needle.
Volume: The volume injected SQ into an adult rat
should not exceed 10 ml.
This route is frequently used as an alternative to intramuscular
injections in the rat. The site usually chosen is the loose skin between
the shoulder blades. Alternatively, the ventral abdomen is commonly used,
employing one handed restraint (Figure
5). The needle is inserted through the skin and advanced 5 to 10 mm
through the subcutaneous tissue to prevent leakage from the site. Rat skin
tends to be thick and difficult to penetrate. Care should be taken to
avoid accidental human injections.
Equipment: Syringe and < 21 g, 1/2-inch needle.
Volume: The volume injected intramuscular in the adult
rat should not exceed 0.3 ml.
This route is usually not used and is not recommended because of the
small muscle mass available and the danger of damaging vital structures.
However, when it is used, the back and hind leg muscles (quadricep;
posterior thigh) are the usual sites selected.
The amount of blood needed and other factors will govern the method and
sites of collection. Descriptions of the various techniques for venipuncture in
different species is available in the Office of the
Campus Veterinarian in text and videotape format. Proper
insertion of the needle into a vein or other part of the vascular system
is normally the most difficult part of the procedure. Certain guidelines
can be given, but only practice provides proficiency. Veins may be
expected to roll, collapse, or shift, making entrance difficult. A
precise, careful introduction of the needle is best and several attempts
may be required. Starting at distal sites will allow repeat attempts more
proximally. The needle is inserted parallel to the vein and the tip
directed into the lumen along the longitudinal axis. When withdrawing
blood from a vein, aspiration should be slow so the vessel does not
collapse.
The area of injection or incision should be cleaned with alcohol. Some
procedures will require sedation or anesthesia; others may be carried out
without anesthesia provided suitable restraint is used. In order to better
visualize veins dilation can be accomplished by immersing the tail in warm
water for 5 to 10 seconds or by warming the animal with a low-wattage
light bulb for 5 to 15 minutes prior to venipuncture. This also aids by
providing additional light.
Equipment: scalpel blade, 25 to 30 gauge needle
The rat is restrained (Figure
6, Figure 7) using
a mechanical device. The veins may be seen laterally near the base of the
tail but good illumination and dilation will normally be required. Dipping
the tail in warm water will help dilate the vessels. A small blood sample
may be collected by capillary action using a microhematocrit tube inserted
into the hub of a small needle previously placed into the tail vein
(future picture). Blood pooling can be induced by placing a small rubber
band around the base of the tail.
Equipment: 22 gauge needle, 3 cc syringe
The rat should be anesthetized and placed in dorsal recumbency.
Bleeding from the tail may be increased by warming it in water at 40C to
50C. Induce arterial dilatation by applying pressure 1 to 2 inches from
the tip of the tail with a finger. Remove the syringe plunger and place
the needle bevel up into the tail artery entering at a 20 to 30 degree
angle (future picture)6.
If placed properly the syringe will immediately begin to fill with blood.
If blood flow is slow or stops, slowly withdraw the needle to re-establish
flow. Retries should always be done with a new needle in a more proximal
location on the tail. Pressure on the site may be necessary to cease blood
flow after needle withdrawal.
Toe clipping or tail
clipping to obtain blood samples: Clipping toes is an unacceptable
method of blood collection. Tail clipping is not a preferable method for
blood collection.
Equipment: 0.90 to 0.50 mm needle
Cardiac puncture represents an accepted method of blood collection from
rats when more than a few drops are required. However, this method also
carries considerable risk to the animal and occasionally deaths occur.
Therefore, it is not recommended as a repetitive blood sampling procedure.
Animals must be anesthetized and restrained in dorsal recumbency. The
needle is inserted under the xyphoid cartilage slightly to the left of
midline (Figure 8). The
needle is advanced at a 20 to 30 degree angle from the horizontal axis to
the sternum to enter the heart. Aspirate lightly while advancing. Blood
should be withdrawn slowly, and the amount must be limited (up to 4 ml in
an adult rat) unless euthanasia is planned.
Equipment: Capillary tubes
Blood collection from the orbital sinus of rats is frequently used.
Bleeding requires that the tube be directed into the orbital sinus (future
picture) which surrounds the globe. In the rat, the capillary tube is
inserted in the medial canthus with gentle rotation while directing the
tube caudally and towards the midline (Figure
9, Figure 10).
Knowledge of the location of the venous structures of the orbit of the rat
aids in establishing a successful peri-orbital bleeding technique.
Pressure should be applied after blood collection to prevent hematomas.
0.5 ml of blood can be obtained weekly using this method. Anesthesia is
required for all peri-orbital bleeding procedures. |
