1. Collection and handling of blood samples The most important aspect of any haematological study or assessment is the quality of the blood sample. The integrity of the cellular and fluid components can only deteriorate once collected from the circulating blood and it is the responsibility of the collector to ensure the best quality sample is collected and delivered to the laboratory for assessment. No matter how good the standard of the laboratory, it cannot compensate for a poor sample. This chapter outlines practices that will aid in the collection of blood, but ultimately it is skill, experience and care that will ensure consistently good quality samples. Collection from veins Blood is usually collected from the venous component of the circulatory system, but may be harvested from the arterial blood if required for a specific reason (such as to assess arterial oxygen tension). The most accessible vein for venepuncture is dependent on the size and anatomy of the species and includes the jugular vein, lateral caudal vein, cephalic vein, saphenous vein and the femoral vein. The most suitable sites for a range of Australian animals are described later in this chapter, but there are some general points regarding blood collection that apply to all sites. Once an appropriate site for venepuncture has been selected, the first and perhaps most important factor in obtaining a blood sample is adequate restraint of the animal. Sedation or general anaesthesia of the patient provides effective restraint. Physical restraint, such as placing the patient in a sack with only the site for venepuncture (limb or tail) protruding or placing a hood on the patient, may be adequate in some cases, but subtle ‘reactive’ movements, especially in response to the needle entering the skin, may frustrate effective venepuncture and the excitement response mediated by increased catecholamine secretion in the restrained, conscious animal may change the haematological values. When the patient has been adequately restrained and the venepuncture site has been selected, the vein must be visualised, which can be facilitated by the removal of hair over the vein using either clippers or scissors. The skin should then be cleaned using an alcohol solution (e.g. 70% ethanol or 100% methanol), or a detergent followed by an alcohol solution. The alcohol should be allowed to ‘air dry’ prior to the venepuncture because contamination of the sample with alcohol may cause some haemolysis (Tietz, 1994). Povidine iodine should not be used as a cleaning agent, as the residue may interfere with some biochemical assays (Tietz, 1994). Once the skin has been cleaned the collector should avoid touching the immediate area to prevent contamination of the site. Table 1.1 Sites for blood collection for selected Australian mammals
INTRODUCTION
GENERAL PRINCIPLES OF COLLECTION
Species | Site(s) for collection of blood | Equipment |
Platypus | Bill sinus | 23–25g needle or butterfly catheter |
Short-beaked echidna | Bill sinus, jugular v., femoral v. | 23–25g needle or butterfly catheter |
Antechinus/dunnarts | Ventral caudal v., cardiac puncturea, orbital sinus | 25g, capillary tube |
Quolls | Cephalic v., femoral v., jugular v., ventral caudal v. | 22–25g |
Tasmanian devil | Femoral v., cephalic v., jugular v. | 22–23g |
Bandicoots | Femoral v., lateral caudal v. | 22–25g |
Bilby | Jugular v., lateral caudal v., femoral v. | 22–25g |
Wombats | Radial v., cephalic v., medial metatarsal v. | 20–22g |
Koala | Cephalic v., femoral v., jugular v., marginal ear v. | 22g |
Brushtail possums | Jugular v., lateral caudal v., ventral caudal v., ear v. | 22–25g |
Ringtail possums | Marginal ear v., ventral caudal v. | 25g |
Pygmy-possums/small gliders | Caudal tibial artery, jugular v., lateral caudal v., ventral caudal v. | 25g, capillary tubes |
Kangaroos/wallabies | Lateral caudal v., cephalic v., recurrent tarsal v., jugular v. | 20–22g |
Rock-wallabies | Lateral caudal v., cephalic v., recurrent tarsal v., jugular v. | 22g |
Pademelons/small | wallabies Lateral caudal v., cephalic v., recurrent tarsal v., jugular v. | 22–25g |
Flying-foxes | Wing v. | 25g needle/butterfly catheter |
Microchiroptera | Wing v. | 25g; capillary tubes |
Rats/mice | Cardiac puncturea, jugular v., lateral caudal v., orbital sinus | 25g, capillary tube |
Dingo | Cephalic v., jugular v. | 20–22g |
Sea-lions/fur-seals | Gluteal v., brachiocephalic v. | 18–20g, 38 mm |
Southern elephant seal | Intervertebral v. | 16g, 77 mm |
Dolphin | Fluke v., dorsal fin v., pectoral fin v. | 18–20g 40 mm, needle/butterfly catheter |
a: Cardiac puncture is not recommended for routine collection of blood.
v.: vein.
The vein should be occluded to impede the flow of venous blood, thus congesting the vessel and facilitating visualisation of the vein. This is achieved by application of a tourniquet, by digital pressure applied by an assistant or by using the non-preferred thumb and/or forefinger of the operator. In larger animals, having an assistant use digital pressure to occlude the vein is the usual technique, but in small species, a rubber band fastened by a haemostat can be used as a tourniquet, which decreases the likelihood of injury to an assistant. Palpation of the area may also aid in identifying the course of the vein; for larger veins, tapping the distal end of the suspected vein should produce a palpable or visible ‘fluid wave’.
Collection using a needle and syringe
In most cases a needle and syringe is used to collect the sample of blood. The size and gauge of the needle and the size and volume of the syringe should be appropriate for the size of the vessel and the volume of blood to be collected. These are usually decided by the size of the animal and a guide for some species is given in Table 1.1. Both the needle and syringe should be sterile. Needles should not be reused between animals because of contamination of samples and possible transmission of disease.
Align the needle, with the bevel facing upwards, and the syringe with the direction of the vein and at an angle of approximately 15° to horizontal. Advance the needle to pierce the skin, subcutaneous tissue and vessel wall. Avoid lateral movement of the tip of the needle as laceration of the vein may result with subsequent haemorrhage and haematoma formation.
Ideally, venepuncture requires only one movement to pierce the vein, but in practice some redirection of the needle may be required to place the needle within the vein. If blood is not obtained after one or two attempts to redirect the needle, withdraw the needle and reassess the situation. In some cases, the difficulty may be the mobility of the vein, which can be limited by the operator placing an extended digit alongside the vein to restrict its lateral movement. Another common problem is when the needle is driven too deeply into the tissue and passes through the vein, in which case the needle should be slowly withdrawn while creating a small amount of negative pressure with the plunger of the syringe until blood appears in the hub of the needle.
Once the needle is aligned within the vein, draw back the plunger of the syringe using gentle, even pressure throughout the length of the syringe until an adequate amount of blood has been collected. Vigorous suction should be avoided as it may cause haemolysis. Excess suction may also cause the vein to collapse, which hinders withdrawal of blood, particularly with small veins. The collector must be patient and apply only very gentle pressure to the plunger of the syringe or use an alternate collection procedure. When the flow of blood from the vein is slow the sample may clot within the syringe, which can be prevented by flushing the needle and syringe with an anticoagulant solution, such as heparin or ethylene-diamine-tetra-acetic acid (EDTA), prior to use (note that EDTA will affect the accurate measurement of some biochemical analytes, such as potassium, calcium and some enzymes).
In many cases the failure to collect a blood sample is because of inadequate restraint of the patient or impatience of the collector (not waiting until the vein is congested and able to be easily visualised). In all cases, venepuncture is made easier by practice.
Once blood has been collected into the syringe, withdraw the needle and apply digital pressure to the vein for a period of 30 seconds to 1 minute to prevent blood flowing from the damaged vein. Remove the needle from the syringe (squirting the blood through the needle will result in haemolysis), gently expel the blood into an appropriate container containing anticoagulant (Plate 1) and carefully mix it. Needles should not be recapped following use, to avoid needle injuries, and both needles and syringes should be disposed of in appropriate biohazard containers.
The blood of some species of Australasian mammals may contain zoonotic organisms. Recently discovered viruses of flying-foxes (Halpin et al., 1999) which cause a fatal disease in humans, are a salient reminder that all samples should be treated as if they contain a harmful organism.
Collection using evacuated blood tubes
Commercially available evacuated blood tubes, commonly known as Vacutainers, are an alternative to using a needle and syringe to collect the blood sample. When the cap of the tube is punctured by the specially designed double-ended needle, already placed in a vein, the negative pressure withdraws blood from the vein into the tube. Evacuated blood tubes may be of use in larger animals, but are inappropriate for use in small animals as the pressure of the vacuum collapses the vein and precludes withdrawal of blood.
Collection using a butterfly catheter
In some circumstances a butterfly catheter may be more suitable for venepuncture than a needle, particularly when animals are not anaesthetised and may move during the procedure. The butterfly catheter provides increased stability and once placed, is less likely to come out of the vein or to lacerate it if the patient moves. There is an increased ‘dead volume’ in butterfly catheters because of the tubing, making them less suitable for small animals and small veins with a slow flow of blood.
Alternative collection sites
Collection from arteries
Some situations may dictate that an arterial sample of blood be collected; for example, an assessment of blood gas concentration. The mechanics of collecting blood from arteries are similar to those for venepuncture. After collection, apply pressure for a longer period of time following withdrawal of the needle, as the blood pressure of arteries is higher than in veins and significant haematoma formation may result from leakage.
Arteries may be used for routine collection of blood samples, such as the cranial tibial artery in small possums, but in some cases sampling from an artery will be accidental. In some sites the artery may be anatomically close to the vein, for example, the femoral artery is close to the femoral vein, and the collector should be suspicious that the sample of blood is arterial rather than venous when the blood is a noticeably brighter red colour and when the collection is more rapid (because of the higher pressure of the arterial system). It is also possible to puncture both vessels when the vein and artery are in close proximity, in which case a ‘mixed’ (arteriovenous) sample will be collected (Tvedten et al., 2000).
Collection from the heart
Cardiac puncture may be used to collect relatively large volumes of blood from virtually any species, but has potentially untoward sequelae including pulmonary haemorrhage, haemorrhage into the pericardial sac and cardiac tamponade, and fibrosis of the cardiac muscle. These may present clinically as respiratory distress, cardiac insufficiency or sudden death and consequently, cardiac puncture is not recommended for routine collection of blood samples. General anaesthesia is mandatory, for both technical and welfare reasons.
This procedure requires the use of longer needles than would usually be required for superficial veins, and the length and gauge will vary with the size of the patient. Place the anaesthetised patient on its side (lateral recumbency) and palpate the lower section of the thorax for the strongest heartbeat. When the animal’s front leg is in a neutral position over the thorax, the point of the elbow is usually over the required area. A needle (with syringe attached) is used to pierce the wall of the thorax through the intercostal space near the costochondral junction and advanced into the heart. Take care to avoid lateral movement of the needle as this may cause laceration of cardiac muscle or pulmonary tissue and consequently result in haemorrhage. Blood is withdrawn into the syringe and then handled as previously described. Mesothelial cells may be inadvertently harvested from the pericardium during cardiac puncture and subsequently observed in blood films (Plate 1).
Collection from peripheral veins and capillary beds
In many species a small volume of blood may be obtained by rupturing a small peripheral vein, such as an ear vein or the lateral caudal vein, using a scalpel or the point of a needle. The drop of blood that wells up into the hub of the needle may be collected into a capillary tube. Similarly, a blood sample may be obtained from a closely clipped toenail, which is a site more commonly used in birds than in mammals but has been used in some laboratory animals, including ferrets (Smith et al., 1994). The toe is cleaned with alcohol and then the distal nail is transected at the level of the supporting dermis. The blood that oozes from the disrupted vessels is collected into a capillary tube. Following collection of blood, haemostasis is effected by the application of pressure or topical agents, such as silver nitrate. If the bone is damaged during the procedure then osteoblasts and osteoclasts may be observed in the blood sample (Clark and Tvedten, 1999). Transection of the distal extremity of the tail has also been used in laboratory animals to obtain samples of blood (Smith et al., 1994). These two methods are not recommended as there are usually alternate methods that allow a greater volume of blood to be collected with less tissue damage.
Capillary blood typically has a lesser haematocrit, haemoglobin concentration and erythrocyte concentration and greater platelet concentration than venous blood (Dacie and Lewis, 1975). Consequently, reference values established for venous blood should not be used for comparison of the results of laboratory analysis of blood collected from capillary beds. Blood from capillaries has been recommended for the investigation of some intraerythrocytic haemoparasites because infected cells tend to accumulate in capillary beds (Jain, 1986).
Collection from the orbital sinus
The orbital sinus has been used as a site to collect blood from small dasyurids and murids, according to Riley (1960).
The donor animal is held by the back of the neck with the left hand, and the loose skin of the head is tightened with the thumb and middle finger. With the aid of the index finger the eye is made to bulge slightly by further traction of the skin adjacent to the eye. The tip of the pipette is then placed at the lower inner corner of the eye and gently but firmly slid alongside the eyeball to the ophthalmic venous plexus which lines the back of the orbit. The venous capillaries forming this network are so fragile that they rupture upon contact with the tip of the pipette and resulting hemorrhage fills the orbital cavity, which serves as a useful reservoir. A slight withdrawal of the pipette frees the tip so that the accumulated blood is drawn into the tube by capillary action. The actual bleeding part of the procedure takes about 2 seconds. Residual blood around the eye is swabbed clear with a soft absorbent tissue to avoid clot formation. Bleeding usually stops immediately upon withdrawal of the pipette and reestablishment of normal ocular pressure on the venous network.
Capillary tubes may be substituted for pipettes. This method allows relatively large volumes of blood to be collected frequently, but requires technically skilled operators, may cause haematomas and optic nerve damage, and is becoming more controversial (Nahas and Provost, 2002). Bradley et al. (1980) reported that repeated samples may be taken by this method with no untoward sequelae, but blindness in two Melomys spp. was believed to have been the result of blood collection using this method (Kemper et al., 1987). This procedure is not recommended for exophthalmic animals because ocular damage may result (Booth, 1999a).
Special considerations when collecting blood
Collection of blood samples in cold climates
When blood is collected from animals during cold weather, the temperature induced vasoconstriction of peripheral vessels may hamper venepuncture. Directing a local source of heat (such as from a lamp) over the site usually provides enough warmth to promote local vasodilation and enable more effective collection of blood from the vein.
Blood volume and collected sample volume
Blood volume may be affected by a wide range of factors, including species, body type, body size, climate, physiological activity, pregnancy and lactation (Jain, 1986), and is commonly reported as millilitres per kilogram of body weight. Blood volume may be determined by measuring both plasma volume and total erythrocyte volume. The former employs colorimetric methods using dyes such as Evans blue or indocyanine green and the latter uses 51Cr labelling of erythrocytes to determine erythrocyte mass. The technical aspects of these methods are discussed by Jain (1986). Some researchers have also used exsanguinations to determine blood volume (Bryden and Lim, 1969).
The volume of blood has been determined for several species of marsupials including the Tammar wallaby (93.5 mL/kg), kangaroos (red, eastern grey, common wallaroo: 87.5 mL/kg) (Maxwell et al., 1964) and the common brushtail possum (51–63.8 mL/kg) (Dawson and Denny, 1968). The blood volume has also been reported for a number of marine mammals including the southern elephant seal (207 mL/kg) (Bryden and Lim, 1969), Weddell seal (186 mL/kg) (Hurford et al., 1996) and New Zealand sea-lion (158 ± 7 mL/kg) (Costa et al., 1998). The results of many studies performed to determine the blood volumes of domestic and laboratory animals have been compiled (Jain, 1986): dog 77–78 mL/kg, sheep 62–66 mL/kg, horse 88–110 mL/kg, rat 70–82 mL/kg and guinea pig 66–78 mL/kg. When the blood volume is not known for a particular species, 70 mL/kg may be used as a reasonable guide.
The volume that may be safely collected (i.e. without challenging circulatory system homeostatic mechanisms) is dependent on the total blood volume of the animal and therefore the size (body mass) of the particular animal. Clinical signs of hypovolaemic shock occur when blood volume is decreased to 60–70% of ‘normal’ (Jain, 1986). A study of rats found no significant alteration in haematological values (haematocrit and haemoglobin and erythrocyte concentrations) when less than 7.5% of blood volumes was removed, and up to 20% of blood volume could be removed without adverse effects on the welfare of the animal (Nahas and Provost, 2002).
Handling of blood samples
Anticoagulants
Once the sample of blood has been collected it must be expeditiously mixed with an anticoagalant to prevent clotting. Several anticoagulants are commercially available, including EDTA, lithium heparin and sodium citrate. EDTA provides the best preservation of cell morphology and should be the anticoagulant routinely employed for haematology. In heparinised blood samples, leukocytes may aggregate and cells stain poorly with Romanowsky stains (compared with EDTA) (Dacie and Lewis, 1975). Consequently, heparin is not recommended as an anticoagulant for routine haematological assays in mammals. Sodium citrate is the anticoagulant that is used when blood samples are collected to investigate haemostatic function. Several sizes of anticoagulant tubes, including 10 mL, 5 mL, 2 mL and 0.5 mL volume (Plate 2), are commercially available and the appropriate sized tube should be selected for the volume of blood collected. Significant underfilling of tubes may result in artefactual changes in the shape of erythrocytes (e.g. echinocytosis).
To minimise haemolysis of the sample, remove the needle from the syringe and gently expel the blood into the tube containing the anticoagulant. Gently roll and/or rock (end to end) the tube so that the blood is thoroughly mixed with the anticoagulant. Vigorous shaking may cause haemolysis and should be avoided. Take care with small (0.5 mL, ‘paediatric’) tubes to ensure the blood is mixed with the anticoagulant because it may be held stationary by surface tension despite the movement of the tube.
If the blood is squirted through the needle into a tube, shaken too vigorously, subject to delayed processing or exposed to temperature extremes then the erythrocytes will lyse (i.e. haemolysis), which may result in spurious laboratory data such as decreased haematocrit and increased mean corpuscular haemoglobin concentration. Haemolysis may also interfere with some biochemical assays. Experimental investigation of the effect of haemolysis on biochemical analysis of canine serum samples found that haemolysis consistently interfered with the analysis of creatinine kinase, lactate dehydrogenase, aspartate aminotransferase, lipase, and albumin, all of which increased with increasing haemolysis (O’Neill and Feldman, 1989).
Blood films
There are several methods that may be used to make blood films. Prior to making any blood film the sample must be thoroughly (but gently) mixed to avoid any sedimentation of cells. The most commonly used technique, the ‘wedge’ method, is suitable for most situations (Figure 1.1). Place a microscope slide on a flat surface (such as a bench top) then place a drop of blood (a generous ‘pin-head’ size) towards the end of the slide. Hold a second slide at approximately 45 degrees to the first slide to spread the drop of blood as follows: touch the ‘spreader’ slide to the first slide in front of the drop of blood, reverse the ‘spreader’ slide into the drop of blood, pause momentarily while the blood spreads laterally towards the edges of the slide and then rapidly and smoothly propel the ‘spreader’ slide forward. Some practical points to consider are:
• if the blood drop begins to dry on the slide, it will not spread well;
• if the spreader slide is stopped and ‘lifted’ at the end of the film, a thick band of blood will form at the ‘leading edge’ of the film;
• if the blood film is too thick or ‘runs’ off the edge of the slide, then too much blood has been placed on the slide;
• if the blood film is too ‘thin’ or too ‘short’, then not enough blood has been placed on the slide;
• if the smear lacks width, the operator has not allowed enough time for the blood to spread laterally before beginning the forward motion of the spreader slide;
• clots and foreign material in the sample usually appear as ‘chunks’ near the leading edge of the film.
Finally, the quality of blood films produced improves with practice and it is often beneficial to make several slides per sample then select the finest example for further processing.
Alternative methods to produce a blood film include those that use two slides, a coverslip and a slide, or two coverslips. The ‘two slide method’ is useful in the field when a clean flat surface is not available for the ‘wedge method’. In this method, a drop of blood is placed on a slide (held by the operator), and then a second slide (held at right angles to the first slide) is flatly touched to the drop of blood (with no downward pressure) and gently advanced along the first slide. Similarly, a cover-slip may be used instead of the second slide. Finally, when only very small volumes of blood are available the ‘two coverslip’ method is most appropriate. A drop of blood is placed on a coverslip then a second coverslip placed on top of the first. The blood spreads under the weight of the second coverslip and when the two are drawn apart, two films are produced.
The gross appearance of blood films can vary with operator and method (Plate 3
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