Administration techniques
Injections and blood samples should be performed using aseptic techniques. New, sterile needles should be used for each animal, and the injection site may need to be cleaned with antiseptic solution prior to administration. This is particularly important for intravenous administration. It is important to use equipment appropriate for the species (see Table 7.2), and use the smallest gauge needle possible. This is determined by the size of the animal and the viscosity of the substance, since thick, viscous liquids may not pass through narrow-gauge needles.
Table 7.2 (a) Suggested hypodermic needle sizes for laboratory animals*. (b) Recommended cannula sizes for laboratory animals.
Subcutaneous injections
For most species, subcutaneous injections can be given into the scruff of the neck (Figure 7.1). A fold of skin is lifted using the thumb and first two fingers of one hand, and the needle is passed through the skin at the base of the fold parallel to the body, to avoid penetrating deeper tissues. Subcutaneous injections are rarely painful, unless the substance being injected causes stinging.
In some animals, subcutaneous injections can also be given over the flank. The animal is restrained as for intraperitoneal injection, and the needle inserted under the skin in the middle of the flank (Figure 7.2). This site is sometimes used for injection of tumour cells, since the growing tumour can easily be assessed in this site, or for antibody production in rabbits.
In large animals, fluid can be given under the skin over the ribs. For pigs, small volumes can be injected into the skin behind the ear, or into the fold between the leg and the abdomen. Pigs have much subcutaneous fat, and injections given elsewhere are likely to enter the fat, where absorption is likely to be slow.
Intramuscular injections
Intramuscular injections are frequently painful, due to the distension of muscle fibres, and an alternative route should be used if possible. If not, good restraint or anaesthesia is required. In small animals they are usually given into the muscles of the thigh, and larger volumes should be injected into the quadriceps group on the front of the thigh1. The muscle can be immobilised with one hand while injecting with the other. In rats, the quadriceps can be palpated on the front of the thigh, and immobilised with the thumb and forefinger of one hand while injecting with the other (Figure 7.3). Injections can be given into the caudal thigh muscles, but as the sciatic nerve runs through these muscles care must be taken here or damage may be caused to the nerve. The risk can be minimised by inserting the needle into the muscle mass from the lateral aspect and angling the point caudally.
In mice, because the muscle masses are so small, a small needle (29 or 30 G) is required and only a small volume can be injected. Injections can also be made into the anterior tibial muscle on the anterolateral aspect of the lower hind limb. The mouse is held on its back with the leg pulled forward. It is important that the tibial tuberosity, a ridge on the front of the leg, is vertical. The needle is inserted at a 45° angle about 3 mm lateral to the tuberosity, parallel to the line of the leg. The needle is inserted to a depth of 2–3 mm. Up to 50 μl can be injected5.
In dogs, cats, ferrets and rabbits, injections can be given with care into the muscles on each side of the spine, and in large animals the gluteal muscles over the rump are used. In adult pigs, injections are given into the neck muscles, but a long needle is required to penetrate the fat layer. Piglets can be injected by lifting them by one hindleg and injecting into the caudal thigh muscle on that side. Fowl are given intramuscular injections into the pectoral muscles. After the injection, the site should be massaged to disperse the dose.
Intravenous injection
In rats and mice, intravenous injections are usually given into the lateral tail veins (Figure 7.4a). Injection is facilitated by warming the animal in a box at 37°C or placing the tail in warm water before injection. Local or general anaesthesia may be needed. The animal should be restrained by an assistant or placed in a restraining device, and the vein can be raised by gently occluding it at the tail base. The needle should be aligned parallel to the vein pointing towards the body with the bevel uppermost before sliding it gently into the vein. There should be no resistance to injection. This method can be seen at www.procedureswithcare.org.uk. In small animals injections can also be given into the saphenous vein (Figure 7.4b). The hindlimb is held in extension, and the vein raised by occluding it at the stifle joint. This is particularly useful for tail-less rodents.
FIGURE 7.4 (a) Tail vein injection in the rodent. (b) Saphenous vein. Photos: (a) MRC Harwell; (b) AstraZeneca, used with permission.
In dogs, cats, primates and ferrets the cephalic vein on the anterior aspect of the forelimb can be used. The vein is raised by placing the first two fingers of one hand behind the elbow and applying pressure with the thumb just below the anterior aspect of the elbow: in ferrets, a tourniquet is best for this purpose as the limbs are short (see Figure 7.5). The jugular vein can be used in dogs, cats, hamsters and ferrets (see Figure 7.6), and this is the method of choice in ruminants and horses. The ear veins are used in guinea pigs and pigs. Fowl can be injected via the brachial vein.
FIGURE 7.5 (a) Quick-release tourniquet. (b) Quick release tourniquet in use for cephalic venepuncture in a macaque.
FIGURE 7.6 Jugular venipuncture in a ferret. Note the needle has been bent at an angle to the hub to facilitate location in the vein.
If fluids are to be administered by infusion, the flow rate should be as low as possible, and the infusion given over as short a time as possible. Otherwise, if too much fluid is given too rapidly, the circulation may become overloaded, causing pulmonary oedema. The maintenance requirement for fluid is approximately 50–60 ml/kg per 24 h in normal mammals and care should be taken not to exceed this unless there are deficiencies to address.
Intradermal injections
Intradermal injections for most species can be given in the same area as subcutaneous injections. However, the dermis of most laboratory animals is very thin, and intradermal administration may require very careful needle placement under anaesthesia or sedation. The site should be clipped to remove fur and cleaned with antiseptic solution to facilitate visualisation of the injection site. The needle should be inserted bevel uppermost a short distance into the skin and the injection given. There will be resistance to injection, and successful administration should result in a raised bleb. The volume which can be injected in this site is small.
Footpad and tail-base injections
The footpad is a commonly used injection site for studying immunological phenomena in rodents. Footpad injection is a combination of intradermal and subcutaneous injection. After injection there is a delayed-type hypersensitivity reaction, which can be used to measure immune responses in vivo. The lymph drainage from footpad injections goes firstly to the popliteal lymph node6, and then to the medial iliac7 and inguinal nodes8. Lymphocytes can be harvested from these nodes for study in vitro. This method of administration is also used in the popliteal lymph node assay, to distinguish between immunostimulating and inert chemicals9,10. Foot-pad immunisation followed by boosting at the base of the tail is also a commonly used protocol for production of antibodies, or for induction of arthritis. However, footpad injection results in swelling and pain and can impair mobility. A more humane alternative to this method is to immunise animals in the lateral tarsal region just above the ankle. The immune response from this site is directed to the same draining lymph nodes, with a similar response, but there is minimal impact on mobility11. Following footpad injections animals must be given soft bedding, and may be provided with food and water in a gel placed on the floor of the cage to avoid the necessity for them to walk around. Social animals should be kept in groups but at a low stocking density so they do not have to climb over each other to reach food or water.
Intraperitoneal injections
Intraperitoneal injections in rodents can be given into the caudal left or right quadrant of the ventral abdomen, to avoid the vital organs. The quadrants are demarcated by the midline and a line perpendicular to it passing through the umbilicus. In mice, both quadrants can be used, although in rats the animal’s right quadrant may be preferred as the larger caecum can occupy the left quadrant. The animal should be held either by an assistant, or in one hand on its back, upright, so that it is comfortable and securely supported. The needle is inserted into the centre of one of the quadrants, angled towards the middle of the back. The needle is inserted a few millimetres, just enough to penetrate the body wall, and the injection given. There is no resistance to injection (Figure 7.7).
Transdermal administration
Some compounds can be introduced into the epidermis by transdermal administration. Particles can be coated onto gold particles, or liquids coupled to drug-delivery systems and fired through the skin using a gas gun. This is often used for vaccination with naked DNA in immunology studies. The advantages of this technique are that it is painless and good immunological responses can be obtained with small quantities of DNA. However, it does require specialised equipment and the specialised preparation and storage of the compound5. Depending on the species and site of administration, light anaesthesia or sedation may be required to allow site preparation and/or administration of the test material. Animals with dense fur such as rodents and rabbits should have the administration site clipped and shaved to remove as much hair as possible. The device or delivery system should be positioned so that it is in close contact with the skin, usually perpendicular to the skin surface. Following delivery the site can be monitored for dermal responses including erythema and oedema.
Oral administration
Substances may be given orally by inclusion in the diet or drinking water. These have the disadvantages that either a whole group has to be dosed, or animals have to be singly housed. Also it is impossible to be sure that the animal has had the entire dose, and some animals may refuse to eat and drink if the feed or water is medicated. In mice particularly, adding drugs to the water can lead to dehydration, because avoidance of drinking by a mouse can lead to a rapid deterioration in the animal’s condition. With ad libitum feeding, or if there is an increase in metabolic rate, the animal may overeat and thus ingest an overdose of the drug. If the watering system is automated it is impossible to give compounds in this way. To overcome these problems gastric intubation or gavage may be employed (Figure 7.8). Flexible catheters or stainless steel needles with rounded tips are used. Rigid needles may be straight or curved. The needle or catheter should first be measured against the animal, and marked at the length which reaches the xiphisternum. The animal is restrained with its neck extended, and the needle or catheter is passed in the midline at the top of the mouth and over the back of the tongue gently down the oesophagus, until the mark on the catheter is reached. Care must be taken not to damage the oesophagus, or to put the needle into the trachea. There should be minimal resistance as the catheter passes down the oesophagus, although there may be some resistance as the catheter passes through the sphincter into the stomach. The needle or catheter can usually be observed passing down the oesophagus on the left side of the neck. Damage to the catheter from chewing can be avoided by using an oral speculum, or by using a flexible nasogastric or pharyngostomy tube instead.
For larger animals, such as dogs and cats, tablets may be administered orally in the conscious animal by placing one hand over the top of the head, placing the thumb at the commissure of the lips on one side and the fingers at the other, and tilting the head back. This will cause the mouth to open slightly. The tablet can be held between the thumb and forefinger of the other hand, and the middle finger used to open the mouth. The tablet can then be placed onto the tongue as far back as possible, to stimulate the swallowing reflex. This can be facilitated by holding the animal’s mouth closed, tilting the head back and massaging the throat.
Legal considerations12,13
The UK Veterinary Medicines Regulations cover the administration of veterinary medicines to animals. These regulations are remade annually and therefore change slightly from year to year. The Veterinary Medicines Regulations 2011 came into force on 1 October of that year. The regulations prevent the supply of medicines for use in animals without an appropriate marketing authorisation, except in particular circumstances. Marketing authorisations do not generally cover laboratory species; therefore medicines used in these species are usually prescribed under an exemption. Supply of medicines for use in laboratory animals is primarily the responsibility of the designated veterinary surgeon, and researchers will usually have to obtain and use these medicines under the direction of a veterinary surgeon. A pharmacist may only supply licensed medicines for use in animals under the direction or prescription of a veterinary surgeon. Medical practitioners may not legally prescribe medicines for use in animals. It is an offence to supply out-of-date drugs.
The regulations categorise veterinary medicine according to the following.
- Prescription Only Medicine–Veterinarian (POM-V): these drugs may only be supplied or prescribed by a veterinary surgeon following a clinical assessment of the animal, or group of animals, under the veterinary surgeon’s care.
- Prescription Only Medicine–Veterinarian, Pharmacist, Suitably Qualified Person (POM-VPS): these may be prescribed by any registered qualified person (RQP; a veterinarian, a pharmacist or suitably qualified person, or SQP). A clinical assessment of the animal is not required, but sufficient information about the animal must be known to prescribe appropriately.
- Non-Food Animal–Veterinarian, Pharmacist, Suitably Qualified Person (NFA-VPS): these may be supplied by an RQP as above provided the requirements for supply are met. These medicines do not require a prescription.
- Authorised Veterinary Medicine–General Sales List (AVM-GSL): there are no legal restrictions for the retail supply of veterinary medicines in this category, but a responsible approach to supply is expected.
Some drugs are also controlled under the UK Misuse of Drugs Regulations 200114. These regulations place additional controls on the supply and use of drugs which are potentially addictive or dangerous. Drugs are classified according to five schedules, each with different requirements for supply, storage and record keeping. Veterinary medicines which are controlled drugs are in schedules 2, 3 or 4. Examples of schedule 2 controlled drugs include morphine and fentanyl. These drugs have to be kept in a locked cabinet, and acquisitions and individual uses recorded. Schedule 3 drugs include buprenorphine. These have to be kept in a locked cupboard but individual administrations do not have to be recorded. Schedule 4 drugs include ketamine and many benzodiazepines, and these must be kept securely and acquisitions recorded. There are strict requirements for the form controlled drugs registers have to take. See reference 14 for details.
Storage and record keeping
All drugs must be stored appropriately to ensure that they maintain their full activity. Some must be kept in a refrigerator at 2–5°C, some need to be kept away from light, some should not be kept in plastic bottles. The data sheet supplied with the drug will specify the requirements. It is good practice to keep a record of all drugs held and used, and it is a legal requirement to keep records of purchase and use of all controlled drugs listed in Schedules 1–3 of the Misuse of Drugs Regulations 200114. It is recommended that an annual check of the drug storage cupboard be carried out and any out-of-date drugs disposed, in accordance with any legal requirements.
Removal of Blood
The removal of blood from an animal has considerable potential to cause pain, suffering, distress or lasting harm. Possible adverse effects are listed below.
- Distress from handling and restraint: humane methods of handling and restraint must be used (see species chapters), and sedatives or anaesthetics may be needed. Training the animals to accept the handling required to take blood samples may also be beneficial.
- Pain and discomfort: venipuncture requires considerable skill, and if carried out incompetently can lead to haemorrhage, bruising, thrombosis, embolism or phlebitis. In addition, poor sampling technique may cause the sample to clot or haemolyse, rendering results invalid.
- Hypovolaemia: the effects of removal of blood depend on the volume of blood removed, and the speed of withdrawal. The rapid removal of large quantities of blood may cause the animal to go into hypovolaemic shock, and may even cause death. The percentage blood loss required to cause hypovolaemic shock varies with the speed of withdrawal, whether or not fluid is replaced concurrently, and the physiological state of the animal at the time. Chronic slow haemorrhage is tolerated better than acute blood loss, and placid animals tolerate greater losses than nervous ones, reiterating the need for competent handling and training of the animals.
Stress responses in the animal result in the release of hormones and other substances to counteract the stress, which can cause anomalous experimental results. Expertise must be gained first by watching others, then by practising on cadavers or models, and then by carrying out the technique under direct supervision. The experimental technique should be refined such that the quantity of blood removed is minimised. This is particularly important in small mammals, such as mice, where the blood volume is small and sample volume is critical. It is essential to follow good practice guidelines such as the BVA/FRAME/RSPCA/UFAW (1993) working group report on the removal of blood from laboratory mammals and birds15.
Quality of samples
To achieve meaningful results, samples must be of good quality, and be preserved in the best possible manner. Samples must be taken skillfully, and treated appropriately thereafter. Blood may be collected using syringes and hypodermic or butterfly needles, through indwelling cannulae, with double-ended needles and evacuated tubes (e.g. Vacutainers, Becton Dickinson), or in very small species by careful incision of a vein using a sterile lancet or scalpel blade and collection of the sample with a capillary tube. If needles are used, the needle should be as large as is practicable for the species. This allows blood to flow faster, reducing the likelihood of clotting, and also causes less damage to the red cells, reducing the possibility of haemolysis.
If no anticoagulant is used, the blood will clot, and serum can be removed after centrifugation. Alternatively blood may be collected into an anticoagulant. Different anticlotting agents are available, including the following.
- Lithium heparin: this is the anticoagulant of choice for most biochemical assays. The yield of plasma from heparinised blood is greater than the yield of serum from clotted blood, which may make heparin a good choice for collecting blood for harvesting antibodies. Sodium heparin is sometimes used if preservation of the white cells is required.
- Potassium ethylene diamine tetra-acetic acid (EDTA): this is used for haematological analyses as it preserves the cells.
- Oxalate/fluoride: this is used for blood glucose determination.
Several other anticoagulants are available, for example for collecting blood for transfusions or analysis of clotting factors. After collection into anticoagulant, the blood should be mixed thoroughly by rolling, not by shaking, as this can damage the cells and lead to haemolysis.
It is preferable for samples to be submitted fresh for analysis. If this is impossible, samples may need to be refrigerated, or deep-frozen after separation of cells and plasma or serum. It is advisable to determine the exact requirements of the laboratory protocol prior to sample collection.
Technique for venipuncture
Site and location of the vein
It is important to be certain of the location of the vein, either by visualising it or palpating its course, and to have it immobilised, before piercing the skin. If unsure of the position of the vein, venipuncture should not be attempted. Sometimes it is necessary to raise the vein, by occluding the venous drainage proximal to the site of venipuncture. This must be performed correctly, or withdrawal of blood will be difficult. The use of a quick-release tourniquet (Figure 7.5a) facilitates blood sampling in some species, and vasodilating agents may be used in others. Warming the animal in a thermostatically controlled hot box for 30 min at 37°C or dipping the tail in warm water can help dilate the tail veins in small rodents, facilitating venipuncture.
Preparation of the site
Blood should be collected using aseptic technique. The area should be clipped to remove hair if necessary, and then cleaned using a suitable antibacterial cleanser such as chlorhexidine. Iodine-based preparations may also be used but are less persistent. The use of warm water with or without disinfectant will help dilate superficial veins as well as cleansing the skin. After cleansing, the skin can be swabbed with 70% ethanol or disinfectant, which helps make the vein stand out as well as disinfecting the skin. It may be advantageous to apply local anaesthetic cream (e.g. EMLA, AstraZeneca) to the site 30–60 min before venipuncture to prevent any discomfort.
Taking the sample
For needle venipuncture, the needle should be held bevel uppermost and directed through the skin following the course of the vein. Once in the lumen of the vein, as determined by the presence of blood in the needle hub, the needle should be advanced parallel to the skin up to the hub, so that the body of the needle is in the lumen of the vein. The needle may be bent at an angle, if required, to facilitate location in the vein; for example, in the jugular vein of the ferret (see Figure 7.6).
A similar method can be used for butterfly needles. Over-the-needle cannulae can be used in almost all species, except possibly mice. Choose the largest, longest cannula that will pass into the vein. Clip the fur from a wide area over the vein, and clean the skin as above. Raise and immobilise the vein (an assistant may be required for this), remove the bung from the stylet which runs through the cannula and insert it through the skin until the tip is in the lumen of the vessel and blood is seen in the hub of the stylet. For thick-skinned animals it may be worth making a tiny skin incision first, using the bevelled edge of a needle or a no. 11 scalpel blade, to facilitate passing the cannula through the skin. The cannula can then be advanced into the vein while retaining the stylet: this prevents the stylet from lacerating the vein as it advances up the vessel, and also the stylet blocks the lumen of the cannula so blood may only be withdrawn once the stylet is removed. Flexible cannulae are less traumatic to the tissues than needles, and are therefore less painful to insert and more suitable for long-term cannulation.
Veins will collapse around the needle if attempts are made to withdraw the blood too quickly, so patience is required, particularly with small mammals. For these species, blood can be allowed to drip from a needle or flexible cannula placed in the vein. A syringe may be attached and gentle suction applied for larger animals. Evacuated tubes can be used if the vein diameter allows it. These are quick and easy to use, but can increase the likelihood of haemolysis as blood cells can become damaged by the rapid passage into the low-pressure container. After blood collection, a swab should be applied to the site and the needle removed, while pressure is applied to the vein for 30–60 s to prevent haemorrhage. The site should then be checked to ensure bleeding has stopped and further pressure applied if necessary.
Potential sequelae
Haemorrhage may occur from the punctured vein. If this occurs, pressure should be applied until the haemorrhage ceases. Dressings are available which will accelerate haemostasis (e.g. Kaltostat, Convatec). Bruising may occur if the vein bleeds under the skin. Pressure should be applied as above, and the site rechecked after 30 min. If the bruised area continues to spread, advice should be sought from a veterinary surgeon. The occurrence of thrombosis or phlebitis following venipuncture indicates poor technique, and the method should be reviewed and advice sought from a veterinary surgeon.
Sample volume
The volume of the sample taken is determined by the requirements of the experiment, and by the safe limit which can be withdrawn without causing distress to the animal. In general, as small a volume as possible should be taken. As a general rule, for mild severity not more than 10% of the blood volume should be removed at one time, and less than 15% of the blood volume should be removed in any 30 day period. Animals have approximately 70 ml of blood per kilogram of body weight, but this varies with the species (see Table 7.3), and withdrawal of a smaller volume may have detrimental effects if the animal is compromised. Larger volumes may be withdrawn with few effects if there is concurrent volume replacement. Maximum limits of volume and frequency of sampling will normally be stipulated in the experimental protocol.
Table 7.3 (a) Practical blood-sampling volumes for laboratory species. (b) Practical blood-sampling volumes for larger domestic species.
Methods of venipuncture in different species
Rodents
In rodents with tails, the usual site for venipuncture is the tail vein. The lateral tail veins are usually used as they are larger than the dorsal and ventral veins. Warming the tail first increases the blood flow to the site and makes sample collection easier. Blood samples may be obtained by incising the skin over the vein using a sterile lancet and collecting the blood into a plain or anticoagulant-coated capillary tube, or allowing it to drip into a container. Scalpel blades are not recommended as they can easily slip, causing damage to the tail. Larger samples can be obtained from rats by placing a 23–26 G flexible cannula or butterfly needle into the tail vein and allowing blood to drip into a collection pot or applying gentle suction with a syringe. If a butterfly needle is used, the plastic tube should be shortened to prevent clotting within it.
For mice and tail-less animals, the saphenous vein can be used16 (see also http://film.oslovet.veths.no/). The vein is punctured using 23 or 25 G needle, a drop of blood forms at the puncture site and can be collected into a microcapillary tube. The scab that forms at the site can be gently rubbed off to enable serial samples to be collected from the same puncture site. In larger animals this vein can be cannulated17.
Amputation of the tail tip to obtain a sample of mixed arterial and venous blood can be carried out on rats, mice and gerbils, but must be done under general anaesthetic21. In mice, this does not appear to involve the removal of any vertebrae, which it does in rats. This should only be performed on a single occasion.
Large samples can be obtained from rats and gerbils via jugular venipuncture, usually under general anaesthesia. The vein is raised on one side of the neck by applying pressure at the thoracic inlet, and a needle placed through the skin and into the vein pointing towards the head.
Blood can also be obtained by puncturing the superficial temporal vein on the face using a sterile lancet18. The facial veins run up from the neck and divide at the angle of the jaw into the superficial temporal vein which runs to the eye, and the facial vein which runs along the jaw line. Hold the mouse by a large pinch of the scruff of its neck on the left side: the eyes should be bulging slightly. The vein can be punctured as it runs along the lower jaw, directly below the lateral canthus of the eye. In white mice there is a grey dot marking the sebaceous gland on the jaw, and the puncture site is found by moving one eye length back and one eye width up from this gland. In black mice there is a black dot marking the gland. The vein should be punctured boldly with the lancet, and the blood should be allowed to fall into the collecting vessel to avoid haemolysis. Blood flows freely from this site if carried out correctly and care must be taken not to take an excess of blood. This technique requires training and practise before use and anaesthesia is advised (see reference 19).
For some animals with no tails, such as guinea pigs, tiny samples can be taken from the ear veins.
For terminal sampling it is acceptable to perform cardiac puncture, which must be done under terminal general anaesthesia. There are many potentially harmful sequelae to this procedure, such as cardiac tamponade. The heart may be reached by placing the animal on its right side and piercing the left ventricle through the chest wall at the sixth intercostal space, one third of the way up, to obtain arterial blood, or by piercing the right ventricle with the animal on its left side for venous blood. Alternatively, the animal may be placed on its back, and the heart reached by passing the needle under the sternum and through the diaphragm.
Rabbits
Blood can be collected relatively easily from the marginal ear vein using an over-the-needle cannula or butterfly needle. The skin over the vein should be clipped and cleaned using a suitable disinfectant, then 30–60 min before venipuncture local anaesthetic cream should be applied. A peripheral vasodilator may also be applied, 5–10 min before blood collection. Once the vein is engorged, the cannula is inserted and blood can be collected by allowing it to drip into a pot. After collection, the vasodilator is wiped off and pressure applied until the bleeding ceases. Bleeding from the central ear artery is possible, but can result in the formation of large haematomata, which can cause damage to the ear or even necrosis.
Ferrets
For tiny quantities of blood, a toenail can be clipped and a drop of blood collected into a capillary tube. For moderate-sized samples the saphenous or ventral tail vein can be used, while larger quantities can be collected from the jugular vein. The fur on the neck needs to be well clipped, and the vein raised by placing a thumb over the jugular groove in the thoracic inlet. The needle can be inserted pointing up the vein towards the head, or down towards the thoracic inlet. The skin is very thick, and some pressure may be needed to penetrate the skin. Collection can be facilitated by bending the needle to an angle of 30° prior to penetrating the skin, or by inserting the needle through the skin parallel to the vein and then into the vein. Pressure on the vein in the thoracic inlet is maintained until the blood has been collected (see Figure 7.6).
Primates
The best method of blood withdrawal in primates is to use the femoral vein, in the groin. This vein is not readily visible and its position has to be determined by palpation. The needle is inserted in the femoral triangle, slightly medial to the femoral pulse, and directed in a craniomedial direction. The needle should be advanced slowly until the vessel is reached, as determined by blood in the hub of the needle. For larger Old World monkeys the cephalic vein on the top of the foreleg below the elbow can be used, as for cats and dogs. The jugular vein can be used as an alternative route. A microcapillary tube can be used to collect small samples from the heel of primates without anaesthetic after they have been trained to accept minimal restraint. Marmosets may also be bled from the coccygeal vein.
Dogs and cats
Small and medium samples can be collected from the cephalic vein one the foreleg. The skin over the anterior aspect of the foreleg is prepared by clipping and cleaning, then a handler restrains the animal by placing one arm over the animal’s body, holding the leg forward by placing two fingers behind the animal’s elbow, raising the vein with the thumb over the top of the leg and restraining the animal’s hindquarters against the body with the elbow. The other hand is placed under the chin to raise the animal’s the head. They can also be bled from the jugular vein. A handler places their right arm over the body of the animal to hold the forelegs. The elbow is used to hold the body of the animal to the body of the handler (Figure 7.9). The left hand is placed under the chin to raise the head. The person collecting the blood raises the vein by placing a thumb in the jugular groove at the thoracic inlet.
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