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Guidelines for Handling, Restraint, Injections and Blood Collection from Small Laboratory Animals

Mice - Rats - Hamsters - Guinea Pigs - Rabbits

These guidelines have been developed to introduce investigative staff to procedures recommended for handling and restraint of small laboratory animals. In addition, techniques for performing parenteral injections and blood collection are also reviewed. This document is intended to supplement hands-on instruction by an experienced member of your laboratory or a member of the Office of the Campus Veterinarian (OCV) staff. OCV staff members are available to provide hands-on instruction. Training workshops can be organized and scheduled for your laboratory.

There are a variety of other techniques, in addition to those described in this document that are suitable alternatives. You can contact the OCV to discuss their suitability or describe them in your animal care and use protocol. Additionally, the OCV library contains more detailed training materials that are available on request .

Handling and Restraint

Although there are significant species differences when handling and restraining an animal, there are several important concepts that apply equally to all species. These include:

  1. Handle animals gently but firmly.
  2. Approach an animal slowly but purposefully.
  3. Wear disposable gloves whenever possible.
  4. Always wash your hands prior to and after handling as odors of other species or blood is frequently distressing, and your hands can act as a means of spreading infectious agents from one group of animals to another.
  5. Wear a clean laboratory coat for the same reasons as discussed above.
  6. Use an appropriate method.

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.

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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 - 1” 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 0.5” 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 0.5” 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 0.5” 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 1” 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 (1”) 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 Ōover the needle” catheter may be inserted if larger volumes are to be administered or repetitive injections will be given. A 0.5” 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.

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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 1” 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 (1.5”) 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 1” 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 (1.5” >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


Credits

American Association for Laboratory Animal Science
University of Washington Training Series
Harkness JE and Wagner JE, The Biology and Medicine of Rabbits & Rodents.


 
 
                     
                         
                         
 
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