Handling of laboratory mice and Different routes of drug administration Handling of Mice Lift the mouse by grasping the tail at the caudal end with the left hand, and allow it to grip the wire mesh of the cage with its forelegs. Grasp the nape of neck between the thumb and the index finger. Place the tail from the hand to the left small finger so that the mouse is held tightly in the left hand. The mouse is now ready for injection.
Introduction To Routes Of Administration Of Drug The The comm common on rout routes es of admi admini nist stra rati tion on of drug drugss are are oral oral,, subc subcut utan aneo eous us,, intramus intramuscula cular, r, intraveno intravenous, us, intrathe intrathecal, cal, rectal, rectal, nasal, nasal, and topical topical applicat application. ion. Each of these routes has its particular advantages. Some substances may be effective when administered by one route but relatively or completely inert when given by another. The mode of entry of a drug into the body determines, to a large extent, the character, speed, dose and degree of action. A. Oral Admiinisration Admiinisration Administration by a stomach tube or by an oral feeding needle. The mouse is held firmly and the feeding needle is held well towards the back of the mouth and near the upper palate. The needle is than passed-gently-not-forced down the esophagus. As it passes into the stomach (about 2 inches down) the animal will exhibit a definite and characteristic gagging. As much as 0.5m1/10 gm of animal may be administered. administered. B. Intravenous Injection The mouse is to be held firmly by wrapping it in a cloth or by placing it in a suitable animal holder. With a little practice the tail veins can be recognized readil readily. y. It is helpfu helpfull to dilate dilate the veins veins by warmi warming ng before before attemp attemptin ting g the injection. Employ a sharp, 26-gauge needle and slowly inject 0.9% saline,(10 ml/kg of body weight) into the mouse. If the needle is not in the vein, resistance will be felt while injecting and blanching will be observed in the surrounding connective tissue. C. Intraperitoneal Intraperitoneal Injection This is a common method to introduce drugs into animals. The drug is injected intothe peritoneal cavity where absorption downward, inject into the lower half of the abdomen; intraintestinal or intravisceral injection may be avoided. Inject 0.9% saline, (10 ml/kg of body weight) into the mouse. D. Subcutaneous Injection The drug is to be introduced directly underneath underneath the skin. A 3/4 -l inch 26-gauge needle is employed. Insert the needle to its full length to avoid loss of liquid upon withdrawal. Inject sample of interest (0.1 ml/l0 gm of body weight) under the
skin of a male mouse followed by gentle massage of the site of injection. The common site employed is the dorsal neck area and that of the abdomen. E. Intramuscular Injection Inject sample of interest into the gluteus maximus muscle of a mouse, using a 26-gauge needle. One must always pull back on the plunger to make certain the needle is not in a vein.
Techniques of blood collection in laboratory animals Introduction Blood is collected from laboratory animals for various scientific purposes, for example, to study the effects of a test drug on various constituents, such as hormones, substrates, or blood cells. In the field of pharmacokinetics and drug metabolism, blood samples are necessary for analytical determination of the drug and its metabolites. Blood is also needed for some in vitro assays using blood cells or defined plasma protein fractions. The techniques for blood collection depend on specific factors which differ from one experiment to the other. There is a difference between terminal and nonterminal blood collection techniques. The conditions of blood collection at the end of an experiment which includes death of the animal (terminal experiment) are completely different (anesthesia, volume of blood) from those of single or repeated blood collections from a conscious animal. Terminal blood collection under anesthesia allows the use of techniques which are not acceptable for non-terminal blood collections. Volume of blood to be removed The volume of blood removed and the frequency of sampling will be based on the purpose of the scientific procedure and the total blood volume of the animal. For reasons of good animal welfare and science, serious consideration must be given to the combined effect of sample volume and the frequency of sampling. If too much blood is withdrawn too rapidly, or too frequently without replacement, an animal may go into short-term hypovolaemic shock and/or in the longer-term suffer anaemia. Data interpretation and scientific validity may be confounded if excessive sampling is employed. ♦ As a general principle, sample volumes and number of samples should be kept to a minimum. ♦ As a rough guide, up to 10% of the total blood volume can be taken on a single occasion from a normal, healthy animal on an adequate plane of nutrition with minimal adverse effects; this volume may be repeated after 3-4 weeks. For repeat bleeds at shorter intervals, a maximum of 1.0% of an animal's total blood volume can be removed every 24 hours; the effects of stress, site chosen and anaesthetic used, must be carefully considered. ♦ If frequent samples are necessary, the use of cannulation as a less stressful alternative to repeated venepuncture should be considered. ♦ See the decision trees for sampling from the mouse and sampling from the rat.
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As a general rule, total blood volume can generally be estimated as 55 - 70 ml/kg body weight. However, care should be taken in these calculations as the percentage of total blood will be lower (-15%) in obese and older animals.
Aspects of animal welfare Minimizing any pain and distress in laboratory animals during the procedure have to be as important as achieving the desired experimental results. This is important not only for humanitarian reasons but also as part of good scientific practice. Blood collection may be stressful to the animal due to the handling and the discomfort associated with a particular technique. Many biochemical and physiological changes are associated with stress, which affect the results, e.g. increases in the blood levels of catecholamines, prolactin and glucocorticosteroids can influence certain metabolic parameters, such as glucose, as well as the counts of erythrocytes, white cells, and packed cell volume. Therefore, stress should be reduced to an absolute minimum if it is not possible to avoid it at all, this is not only in the interest of animal welfare but also in the interest of good science to obtain representative data. During non-terminal blood collection it is important not to withdraw too much blood which could reduce total blood volume and lead to false results. A reduced total blood volume is accompanied by a reduced hemoglobin content and oxygen transport capacity as well as by a fall in blood pressure and an increase in the concentrations of stress related hormones. The welfare of the individual animal should not be endangered by removal of too large a volume of blood or by too frequent collections. This may be the case more often when small laboratory animals, e.g. mice, gerbils, rats or hamsters are used. In these cases the study protocol should be adapted to use more animals to minimize distress for the individual animal. Technical aspects of blood removal Tail Snip, Tail Nick method and Lateral Tail Vein Sampling: These methods can be used in both rats and mice by cannulating the blood vessel or by nicking it superficially perpendicular to the tail. A common method in mice and rats for collecting up to 0.1 ml capillary blood is to remove the tip of the tail. For a very small sample, i.e., a single drop of blood, snipping of no more than the distal 1-2 mm of the tail can be a viable alternative. With this method, the clot/scab can be gently pulled for repeat, small samples if needed for blood glucose measures, etc. This method is sufficient for multiple blood collections to determine, e.g. blood glucose or total radioactivity after the administration of radio labeled drugs. Sample collection using a needle minimizes contamination of the sample, but is more difficult to perform in the mouse. Sample collection by nicking the vessel is easily performed in both species, but produces a sample of variable quality that may be contaminated with tissue and skin products. Sample quality decreases
with prolonged bleeding times and “milking” of the tail. When the sample is taken too quickly by a syringe, the vein will collapse. After the needle has been withdrawn, continuous pressure should be applied immediately to the puncture site and maintained for at least 30 seconds. The animal should be monitored 15 min later to check for after- bleeding This method is non-traumatic and routinely done without anesthesia, although effective restraint is required. In most cases warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume. Multiple withdrawal of blood samples should not exceed 1% of total blood volume every 24 h (0.6 ml/kg/d). This technique normally recovers a few drops of blood, adequate for hemoglobin, microhematocrit and cell counts. Larger blood samples can be obtained by making a small incision over the vessels 0.5 to 2 cm from the tail base using a scalpel blade. One half to one milliliter of blood can be withdrawn using this method. Anesthesia or sedation should be used. Number of samples: No more than four blood samples should be taken within any 24-hour period. Sample volume:10 ul Adverse effects : • •
Infection <1% Haemorrhage
Retro-orbital bleeding Blood sampling by orbital puncture is a technique routinely applied in most laboratories. The puncture of the orbital venous plexus is often performed in tailless animals, e.g. hamsters. This technique is also used in rats and mice, when larger volumes are required which cannot be obtained from the tail vein. Basically, retro-orbital bleeding should always be performed under anesthesia. Pasteur pipettes, micropipettes or microcapillary tubes are used and pushed with a rotating movement through the conjunctiva laterally, dorsally or medially of the eye to the back wall of the orbit. Sterile capillary tubes and pipettes are recommended for use to help avoid periorbital infection and potential long term damage to the eye. The edges of the tubes should be checked for smoothness to also decrease likeliness of eye damage. However severe side-effects such as retroorbital haematoma with subsequent pressure on the eye cannot be completely excluded. This pressure can damage the optical nerve. The animal may be unable to close its eye. Bleeding from the orbital venous plexus should only be performed with recovery of the animal in
exceptional circumstances when there is no other method available. The technique should be performed only by a well-trained staff and only one eye should be used. Although the procedure may appear to members of the lay community as unduly distressful, the NIH ARAC has determined that in the hands of a skilled technician retro-orbital bleeding is a humane procedure that produces minimal and transient pain/distress. Retro-orbital bleeding can be conducted in awake mice. Due to pain and distress issues retro-orbital sampling in the rat is best conducted under general anesthesia Rapid and large number of mice/rats can be bled within a short period of time. Number of samples: It is recommended that only one sample be taken. Sample volume: Up to 0.2 ml with recovery; Up to 0.5 ml non-recovery. Cardiac puncture In tailless animals such as guinea pigs and hamsters, cardiac puncture under general anesthesia may be the preferred technique. The collection of blood by cardiac puncture has been performed in guinea pigs, gerbils and hamsters. In these species it is difficult to collect blood by alternative methods except retro-orbital bleeding. In general, cardiac puncture should be performed under general anesthesia with atropine as premedication to prevent cardiac arrhythmia. If cardiac puncture is used for a non-terminal blood withdrawal with recovery, the animal has to be separated from other animals until it is fully conscious. It should be carefully watched for adverse effects and sacrificed if found in distress due to complications like bleeding into the pericardium or into the thorax. The approximate circulating blood volume of rodents is 55 to 70 ml/kg of body weight. Of the circulating blood volume, approximately 10% of the total volume can be safely removed every 2 to 4 weeks, and 1% every 24 hours. Number of samples: One Sample volume: Up to 1 ml Conclusion: The given mice were handled the various routes of administration of drugs were observed and practiced. Also blood collection was preformed using the tail snip and retro orbital bleeding bearing in mind the volume constrains for the methods in question.
