Facts sheet 4

Experimental Techniques and
Anaesthesia in the Rat and Mouse
ANZCCART
Facts Sheet
Steven Marshall*, Angela Milligan** and Ray Yates***
*Austin Hospital, Heidelberg, Victoria, 3084 **Walter and Eliza Hall Institute of Medical Research, ***Flinders Medical Centre, Bedford Park, SA, 5042 Introduction
cylinder with a plunger which can be adjusted to the size of the The use of animals in basic biological research and in research animal, or a cone shaped device, which restrains the body but applied to specific purposes has made major contributions to the welfare of man and other animals in the treatment and Chemical restraint in the mouse and rat
prevention of disease. However, progress of experiments and the quality of life of experimental animals is closely related to theability and training of research personnel and animal care staff One to two ml of an inhalant anaesthetic (halothane, handling the animals. A relatively simple procedure, such as methoxyfluorane or isofluorane) is placed on a cotton wool pad blood sampling, can become a traumatic experience for the in a bell jar or screw top glass jar. This method of anaesthesia is animal if performed roughly or incompetently by an unskilled or often used for short term procedures where the animal needs to unsympathetic person. Familiar staff and competent handling be anaesthetized for only a few minutes. When the liquid has can do much to reduce the fear and distress that many animals been placed on the pad in the jar, the lid is replaced, allowing the may otherwise feel. The following notes have been collated by vapour to fill the jar. The mouse or rat is then placed in the jar experienced animal technicians to provide some guidance in the and removed when fully anaesthetized. Should the animal need use of a number of commonly performed techniques.
to be maintained, the application of a nose cone containing a Handling and restraint
small quantity of the anaesthetic can be carried out at regularintervals. It is always desirable to have a grid between the animal It is very important both for the safety and comfort of the animal and the soaked cotton pad, so that the animal does not physically and of its handler that the animal is properly restrained before contact the anaesthetic. This method of anaesthesia should always be carried out in a fume hood and the container should Mice can be picked up by the base or middle third of the tail always allow easy viewing of the animal.
and placed on the cage lid. They can then be restrained by The inhalant anaesthetic can also be delivered by an grasping the loose skin behind the ears with the thumb and fore- anaesthetic machine, in combination with finger, while keeping some tension on the tail. If the skin is into a nose cone or perspex box with scavenging system grasped too loosely or over the shoulders rather than behind the incorporated. This is much safer than the open drop method if ears, the mouse may be able to turn its head and bite. Once the using halothane or isofluorane, as the animals can become mouse has been ‘scruffed’ it can then be picked up and its tail deeply anaesthetized very quickly and the use of a calibrated held between the fourth and little fingers. It can easily be held machine gives far greater control of anaesthetic depth.
securely, allowing administration of injections or palpation ofthe abdomen.
Injectable anaesthetics (see table 1)
Rats should not as a rule be handled with gloves or forceps.
They learn quickly and will soon become accustomed to the Untrained personnel should always have an assistant when investigator’s hand. They should be approached from behind and attempting any techniques, that is, one person restraining and the grasped firmly around the neck by the thumb and fore-finger, other applying the technique. Only fully trained personnel with one fore-leg encircled. The other hand may be used to should attempt to restrain an animal and give an injection A number of mechanical devices can be used to restrain rats and mice and are usually used for giving intravenous (iv)injections via the tail vein. The animal is restrained either in a Anaesthetic - not required. Needle size and gauge - 25mm/23-27 gauge.
Thigh muscles that can easily be injected are the tensor fascia Contents.
lata, biceps femoris, vastus lateralis and semitendinosus. Anyinjection into these sites risks damage to the sciatic nerve. One Handling and restraint
operator should firmly place an index finger and thumb under Inhalant anaesthesia
the chin (mandibles) of the rat, carefully lifting and supportingthe body while the free hand grasps the base of the tail or pelvis.
Injection techniques
Once the animal is securely restrained it can be presented to the Oral dosing
other investigator to administer the injection. The person Blood collection
administering the injection can firmly grasp the paw of either legand gently stretch the limb so that most of the muscles are Rederivation of a colony
extended. The investigator should insert a short, small gauge Reproductive techniques
needle on a parallel plain to the femur into the posterior muscles, Euthanasia
while aspirating the plunger to ensure that blood vessels have notbeen breached. Only small volumes can be injected References
ANZCCART News Vol 7 No 1 March 1994 Insert
measurements in rats. Once the rat is firmly restrained, the Anaesthetic - not required. Needle size gauge - 25mm/23-27 gauge.
needle can be inserted, bevel up, using a 1, 2, or 2.5ml syringe.
The needle must be inserted at a shallow angle. Once in the Rats are restrained as for the i/m injection. Two major methods vessel, a small amount of blood will appear in the syringe. This of administration are practised. First, the rat is slightly tilted so blood must appear before injecting, as it indicates the needle’s that its head is facing towards the floor. This allows the position and ensures the material is injected in the vein. A welt abdominal organs to move toward the thoracic cavity enabling will appear around the vein if the needle is not positioned the second operator to insert the needle laterally to the midline, correctly. If blood samples are required, slowly withdraw the (imagine a quadrant drawn onto the rat’s abdomen, and injecting plunger until the required volume is collected. Applying into the lower squares), thereby avoiding major organs. The excessive force on the plunger will only collapse the vessel.
second method is to line the needle up with an outstretched legas the needle is inserted into the peritoneal cavity. This Anaesthetic - not required. Needle size and gauge - 13 to 25mm/23 to 26 gauge. Volume - maximum 5ml.
Anaesthetic - is required. Needle size and gauge - 25mm/26-27 gauge.
One trained investigator can carry out this technique, providingthe animal can be fully restrained without undue stress.
Rats ideally should be anaesthetized to perform this technique, Carefully remove the animal from its box or cage and gently as the needle insertion requires the animal to be completely still.
place it on a non-slip surface. Once the rat is carefully restrained Only small amounts of fluid should be injected (up to 100µl).
by the investigator, the needle can be inserted into the animal’s Once the animal is anaesthetized the back can be shaved. This is flank, back of the shoulder or down the animal’s back. The the usual site to administer an i/d injection. The closer the administration site will depend on the method of restraint.
shaving, the easier it is to apply this technique. A small gauge Working on a standard height laboratory bench, the animal can needle is inserted into the dermal layers. If the bevel is facing the be gently pressed against the waist of the operator. One person operator it can be seen through the skin after it is inserted. The can rest a hand on the back of the rat with the index finger and needle should not be inserted past the end of the bevel. A small thumb positioned over the neck. A fold of skin can be gently bulge will appear at the site if the injection is successful.
pinched into the shape of a ‘tent’ while the needle is inserted parallel to the animal’s body with free hand. A perpendicular Anaesthetic - not required. Needle size and gauge - 25mm/26-27 gauge.
needle insertion can result in the investigator jabbing a finger, orpushing the needle through the skin on either side of the fold.
Intravenous injection and collection from the rat tail is one of themost common techniques. Various methods have been adopted by With the investigator sitting at a bench, a rat can also be investigators, based upon convenience, skill and requirements.
restrained by placing it between the forearm and waist. In the Lateral tail vein incision, where a vacuum tube is placed over the case of a right handed person the rat will be restrained by the left tail, has been replaced by the use of needles and syringes. Other forearm, facing the head to the left. Using the left hand a fold of intravenous collection and injection techniques include skin can be pinched and the injection given with the right hand.
cannulation or needle insertion into the femoral and/or jugular The use of two operators will cause less stress to the rat, as less veins and needle insertion into the sublingual or the saphenous time will be required to complete the task.
vein. The lateral tail vein bleed or injection is the most commonly used technique. The rat should be placed in a cleancontainer with no food or water on the lid. A heat lamp should Anaesthetic - not required. Needle size and gauge - 50 to 100 be placed approximately 10 to 20 cm from the cage lid to warm the animal, so that the tail vessel will dilate.
Stomach tubes or catheters are occasionally used to orally dose Rats should not be allowed to become too hot and must be rats. However, the most commonly used apparatus is a metal continually monitored. Once the desired heat is reached the rat needle with the sharp end removed - a gavage needle. Gavage must be quickly removed and restrained for the needle insertion.
needles allow the investigator to administer an accurate dose rate Commercial restrainers are available although a quick, cheap or volume into the oesophagus or stomach. They can be 15 to 18 and more comfortable technique for the rat is to roll it into a gauge, with a ball of silver soldered onto the distal end, which medium sized towel (The Duffy Rat Roll). The rat is placed into will allow smooth entry into the mouth, through the posterior a blind fold of the towel, while the rest of the towel is rolled over oropharynx and into the oesophagus. Needles can be straight or and under the rat. This restraining technique can also be used curved and up to 100mm long. These are available when performing i/m injections or taking blood pressure commercially, although many institutes make their own.
Table 1. Injectable anaesthetic agents
Comments
Anaesthetic effects can vary depending on strain, sex andage of animal. Poor analgesia, low margin of safety.
Excellent but short acting anaesthetic. Will need Excellent analgesia, 30-40 min. surgical anaesthetic but a Slight respiratory depression can be reversed with buprenorphine without loss of analgesic potential.
Poor analgesia and marked respiratory depression. Can cause ileus in rats.
Good anaesthesia but solution must be freshly prepared.
Post-operative death or adhesions can be sequelae.
ANZCCART News Vol 7 No 1 March 1994 Insert
This technique can be used by one person, although two Blood collection from the rat
people should be used for wild or sick animals. A single investigator can restrain a rat by firmly placing an index fingerand thumb under the chin, then carefully lifting and supporting Anaesthetic - is required. Needle size and gauge - 25mm to the rat’s body while pressing the animal against his chest. In this position the animal is lightly restrained and reasonably As anaesthetics are involved in the application of this procedure, comfortable. Finally, tilt the rat’s head into a position so that it is only experienced investigators should attempt it. Although the facing the investigator, enabling the gavage needle to be gently tail veins are in a more convenient location for access in the rat, it is difficult to obtain large volumes of blood from the tail.
Injection techniques in the mouse
Intracardiac bleeds allow the investigator to obtain largevolumes with each application on a fortnightly basis. The number of serial bleeds depends on the previous volumes • prior to administering an injection the animal must be removed, number of anaesthetics administered and the general well- adequately restrained, whether by the hands, a restraining being of the rat. Maximum bleed should not exceed 1% of the rat’s device or assistance from another person, body weight. An experienced investigator will obtain the requiredblood volume within one minute, once the rat is anaesthetized.
• use your dexterous hand to inject and restrain with the other, Placing the animal on a warm or protected surface on its • keep the bevel of the needle and graduations on the syringe back, the investigator can, using the index finger and thumb, in line with each other and facing upwards, locate the beating heart through the left side of the chest • use as small a gauge needle as the substance permits to do the (between the fifth and sixth ribs) and insert the appropriately injection. Use a larger gauge if necessary to draw up the sized needle through the intercostal muscles into the left solution, then change to a smaller gauge, ventricle of the heart. Using the thumb or index finger as a guide • don’t put your finger on the plunger until ready to inject.
for the needle ensures that a straight and direct insertion is made.
If the first attempt is not successful, do not partially withdraw the needle and re-insert it at a different angle. Soft tissue damage Keeping the syringe in a horizontal position, insert the needle can occur, causing internal bleeding. The needle must be under the surface of the skin in the lower abdomen. Inject the completely removed from the thoracic cavity and re-inserted.
mouse with as small a volume as possible. The dorsum of the Left ventricular insertion provides a strong flow into the needle.
mouse can also be used as a site of s/c injection, using loose skin Tail clipping is not recommended, as small or large volumes of blood can be obtained using a needle and syringe. A m p u t a t i n gthe tip of the tail causes more pain than necessary whenother less The skin will swell at the injection site immediately, invasive techniques are available to the investigator.
indicating that the injection is not too deep, a common error.
Leave the needle under the skin for a few seconds post-injecting, then remove slowly as this will help prevent leakage. The larger the volume, the greater the likelihood of leakage.
Orbital sinus bleeding in the rat can be carried out in the same Using the same method as for s/c injections, insert the needle manner as the mouse. The rat needs to be fully anaesthetized for under the skin surface, lift the syringe to a vertical position and the procedure. A pasteur pipette or a haematocrit tube can be push down through the abdominal wall into the peritoneal cavity used for the collection, depending on the volume of blood and inject the animal. Watch the length of the needle and the required. The blood vessels behind the eye are harder to angle at which you inject. It may be helpful to tilt the animal’s penetrate in the rat than the mouse and so a little more pressure head downwards so that the internal organs fall away from the in necessary along with the constant rotation of the tube to injection site. It is not uncommon to inject volumes up to one ml by this route, but the amount injected will be determined by the Blood collection from the mouse
Blood parameters can vary markedly, depending on the site of collection. Once a method has been decided, it should not be varied.
Dilate the tail vein by placing the animal under a heat lamp for two to ten minutes prior to injection. A cone shaped device may The mouse should always be anaesthetised unless this technique be used to restrain the animals. Veins are located either side of is performed by an experienced technician. A microhaematocrit the artery (which runs down the centre of the tail) and appear tube is inserted via the lateral (or medial) canthus, at an angle of blue-red in colour when dilated. Keeping in line with the vein, about 30˚ into the venous plexus behind the eye with a twisting insert the needle, which should be no larger than 26 gauge, into motion and the blood allowed to flow through the tube into the the vein about three cm from the tip of the tail. To check that the collection vessel. If larger amounts of blood (up to 0.5ml for a large needle is in the vein, pull back on the plunger for signs of blood.
mouse) are required, it helps if the microhaematocrit tube is coated If there are no signs, remove the needle and try again closer to with an anticoagulant. Bleeding usually stops when the tube is removed.
the tail butt. Another way of checking is to depress the plungerand, if you feel any resistance, try again. However, if the plunger moves freely, continue to inject. The tail vein will collapse if it This is useful for obtaining large amounts of blood. The mouse must be anaesthetised and it is usually a terminal procedure. The Intravenous injection in neonates is sometimes necessary apical beat is palpated and the needle inserted either through the and can be performed using a 30g needle into the anterior facial ribs or via the diaphragm under the xiphoid cartilage and slightly vein at the level of the lateral canthus of the eye, or into the to the left of the midline, at an angle of 20˚ - 30˚ from the sternum.
transverse sinus, which is easily identified on the surface of the Rederivation of a colony via caesarian section and embryo
cranium. Intra-peritoneal injection of neonates is often transfer
performed by inserting the needle in the skin over the thoracic This may be used to obtain mice free of endemic disease or to cavity towards the animal’s umbilicus, over the liver and then eliminate unwanted pathogens from the colony.
into the animal’s abdominal cavity.
The donor female should be euthanased by cervical dislocation.
This is rarely used in the mouse, due to the very small amount of A small transverse incision is made in the skin on the abdomen muscle mass and the fact that rates of absorption of aqueous and the skin peeled away. The mouse is then submerged in 37˚C solutions are similar to the s/c route of administration. A iodine for 4 minutes. Then open the abdomen and carefully maximum volume of 0.05ml into the quadriceps femoris musclegroup is possible.
* (See elsewhere in this issue of ANZCCART Newsfor a report on this technique) ANZCCART News Vol 7 No 1 March 1994 Insert
remove the uterus, dipping it into warm formal saline before Care must be taken to ensure that the animals cannot come in placing on a warming board. Carefully cut along the uterus contact with the dry ice or tip over the beaker.
spilling the neonates from within. Use a sterile linen swab to Halothane, methoxyfluorane or isofluorane can be used for wipe their mouths briefly, before cauterizing the umbilical cord.
euthanasia, either in an anaesthetic chamber where the Move the neonates to a clean area on the board and spend a few anaesthetic is controlled by an anaesthetic machine and more minutes gently wiping them, which also helps to stimulate vaporizer, or a glass jar, where anaesthetic soaked cotton wool breathing. Once they are breathing and pink in colour, they are or gauze is placed in the bottom. When these agents are used it ready to be fostered. To help the foster mother accept the pups it should be in a fume hood or with apparatus which has a is a good idea to put her own pups on top of the foster pups, or scavenging mechanism, to minimise the risk of exposure to the for the foster mother’s pups to urinate on the new litter, which assists with the transfer of smell. Be careful not to confuse the litters.
Sodium pentobarbitone given i/p at the rate of 10 - 15 Plugging
mg/100g body weight produces unconsciousness, followed Set the animals up for mating and make a habit of checking the female early every day for the presence of a vaginal plug. The Staff performing physical methods of euthanasia must be plug consists of coagulated proteins from the seminal fluid and well trained. Cervical dislocation in the mouse is carried out by can usually be seen by placing the female on the cage lid and placing thumb and forefinger behind the skull, holding it firmly lifting the hind legs off the lid by pressing gently just above the and pulling the tail in the direction away from the body, causing tail base to expose the vagina. The plug is a creamy yellow dislocation of the neck. This can be used in young rats up to colour and when gently touched with a small probe or blunt 150g, whilst larger rats can be stunned by a sharp blow to the forceps will feel solid. Certain strains of animal tend to have back of the head. However, this should only be performed by deep implantations and blunt forceps may be used to gently open operators who have had experience in this technique and are the vagina for checking. The plug usually dissolves about 12 competent. Where the animal’s brain needs to be untraumatised, hours after mating. The day that the plug is first noticed is an alternative is decapitation by guillotine. When this method is used, sure handling and speed reduce the stress on the animal.
Progesterone is administered to pregnant mice on the 17th ANZCCART (1993) published a detailed monograph covering this topic.
and 18th day after the plug to prevent the mouse delivering References and further reading
naturally. This is common practice for animals that are being ANZCCART (1993). Euthanasia of animals used for scientific used for caesarians, which would be performed on the 19th day.
purposes. Reilly, J.S. (ed.). ANZCCART, Adelaide.
0.2ml progesterone* is added to 5ml of sterile peanut oil and0.1ml is then injected s/c into the scruff of the neck. Depending AVMA (1993). Report of the AVMA panel on euthanasia. on the gestation period of the strain, the dates of administration JAVMA 202, 229 - 249.
of progesterone should be adjusted accordingly.
Callahan, B.A., Fiorillo, A.P., Hutchinson, K.A. and Keller, *Medroxy progesterone acetate 50mg/ml(Upjohn).
L.S.F. (1992). A comparison of four methods for sterilizingsurgical instruments for rodent surgery. Contemporary Topics Superovulation
in Laboratory Animal Science. 31 (4): 38 (abstract).
A large number of immature ovarian follicles are induced to Cunliffe-Beamer, T.L. (1983). Biomethodology and surgical maturity by an injection of Folligon, a complex glycoprotein techniques. I n : The mouse in biomedical research. Vol. III.
from the serum of pregnant mares. It is marketed in a freeze Foster, H.L., Small, J.D. and Fox, J.G. (eds.). Academic Press, dried form. Make a 5ml volume by adding the solvent provided.
Withdraw 1ml of the solution, add to 7ml of MTPBS** and mix.
Cunliffe-Beamer, T.L. (1990). Surgical techniques. I n : Inject 0.2ml/mouse i/p, then 44-48 hours later, inject the animal Guidelines for the well-being of rodents in research. Guttman, with human chorionic gonadotrophin (APL)***, which has H.N. (ed.). Scientists Center for Animal Welfare, Bethesda, luteinizing hormone activity. Ovulation should occur approximately 12 hours later. To make up the APL, mix 0.2mlof the solution with 3.8ml of MTPBS. Inject 0.2ml i/p per mouse Cunliffe-Beamer, T.L. (1993). Applying principles of aseptic and put the female in with the fertile male studs. The females surgery to rodents. AWIC Newsletter, April-June 1993. 4 (2), 3 - 6.
should be plugged the following morning. Use females between Flecknell, P.A. (1984). The relief of pain in laboratory animals.
3 - 5 weeks of age, but be aware that the optimum age for Lab. Animal 18, 147 - 160.
superovulation can vary a little between strains. A superovulated Flecknell, P.A. (1987). Laboratory animal anaesthesia. An female can produce up to 50 embryos.
introduction for research workers and technicians. A c a d e m i c **mouse toxicity phosphate-buffered saline.
***APL, Ayerst Laboratories, at a dose of 10i.u. per mouse.
Hogan, B., Costantini, F. and Lacy, E. (1986). Manipulating the Pseudo-pregnant foster mothers
mouse embryo - a laboratory manual. Cold Spring HarborLaboratory, USA.
Successful embryo transfer depends on the quality of the hostmaternal environment. Mice are spontaneous ovulaters and can Kraus, A.L. (1979). Research methodology In: The laboratory be made pseudo-pregnant by mating with vasectomized males r a t . Vol. II. research applications. Baker H.J., Lindsey J.R.
during oestrus. They will display the hormonal profile of a and Weishbroth S.H. (eds.). Academic Press, New York. pp. 2 - 28.
normal pregnant female. Mate the recipient females by 9.00am Olfert, C.D., Cross, B.D. and McWilliam, A.A. (eds.) (1993).
on the same day that the superovulated mice are plugged and Guide to the care and use of experimental animals. Vol. I (2nd observe for plugs the following morning. An excellent description ed.). Canadian Council on Animal Care, Ottawa.
and illustration of embryo transfer and vasectomy can be found Poole, T.B. (ed.) (1989). The UFAW handbook on the care and management of laboratory animals. (6th ed.). Longman Euthanasia in rats and mice
Carbon dioxide inhalation is probably the most efficient and Scott, L. (ed.) (1991). Mouse management. A practical guide to aesthetically acceptable method of euthanasia for both the mouse the care of laboratory mice. University of Melbourne.
and rat. The method is quick and most suitable when large Short, D.J., and Woodnott, D.P. (eds.) (1969). The I.A.T. manual numbers of animals need to be killed. The gas is piped into a of laboratory animal practice and techniques. (2nd ed.).
purpose-built chamber or a plastic bag containing the cages of Charles C. Thomas, Springfield, Illinois.
the animals to be killed from a cylinder fitted with a regulatorand flowmeter. Depending on the size of the chamber, allow Tuffery A.A. (ed.) (1987). Laboratory animals. An introduction several minutes for the carbon dioxide to fill the chamber. If for new experimenters. Wiley Interscience, London.
bottled carbon dioxide is unavailable, dry ice can be placed in a Waynforth H.B. and Flecknell, P.A. (1992). Experimental and container of water inside the chamber containing the animals.
surgical techniques in the rat. Academic Press, London.
ANZCCART News Vol 7 No 1 March 1994 Insert

Source: http://www.aacytal.com.ar/experimental.pdf