How To Collect Blood Sample In Animal Edta Needle

ane. Drove and treatment of blood samples

INTRODUCTION

The almost important attribute of any haematological study or assessment is the quality of the blood sample. The integrity of the cellular and fluid components tin can simply deteriorate once collected from the circulating claret and it is the responsibility of the collector to ensure the best quality sample is collected and delivered to the laboratory for cess. No matter how adept the standard of the laboratory, it cannot compensate for a poor sample.

This chapter outlines practices that will assist in the collection of blood, only ultimately information technology is skill, feel and care that will ensure consistently good quality samples.

General PRINCIPLES OF Collection

Collection from veins

Claret is usually collected from the venous component of the circulatory system, only 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 beefcake of the species and includes the jugular vein, lateral caudal vein, cephalic vein, saphenous vein and the femoral vein. The virtually suitable sites for a range of Australian animals are described later in this affiliate, but there are some full general points regarding blood collection that apply to all sites.

Once an advisable site for venepuncture has been selected, the outset and mayhap well-nigh important factor in obtaining a blood sample is adequate restraint of the animal. Sedation or general anaesthesia of the patient provides effective restraint. Concrete 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, merely 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 fairly restrained and the venepuncture site has been selected, the vein must be visualised, which tin can be facilitated past the removal of hair over the vein using either clippers or scissors. The skin should then be cleaned using an alcohol solution (east.yard. 70% ethanol or 100% methanol), or a detergent followed past an alcohol solution. The alcohol should exist allowed to 'air dry' prior to the venepuncture considering contagion of the sample with alcohol may cause some haemolysis (Tietz, 1994). Povidine iodine should non be used as a cleaning agent, as the residuum may interfere with some biochemical assays (Tietz, 1994). Once the skin has been cleaned the collector should avert touching the immediate expanse to prevent contamination of the site.

Table 1.ane Sites for claret collection for selected Australian mammals

Species	Site(s) for drove of blood	Equipment
Platypus	Nib sinus	23–25g needle or butterfly catheter
Short-beaked echidna	Nib sinus, jugular v., femoral v.	23–25g needle or butterfly catheter
Antechinus/dunnarts	Ventral caudal v., cardiac puncture a, orbital sinus	25g, capillary tube
Quolls	Cephalic v., femoral v., jugular 5., ventral caudal 5.	22–25g
Tasmanian devil	Femoral v., cephalic v., jugular 5.	22–23g
Bandicoots	Femoral five., lateral caudal v.	22–25g
Bilby	Jugular v., lateral caudal five., 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 5., lateral caudal v., ventral caudal 5., ear v.	22–25g
Ringtail possums	Marginal ear v., ventral caudal v.	25g
Pygmy-possums/small gliders	Caudal tibial artery, jugular five., lateral caudal 5., ventral caudal v.	25g, capillary tubes
Kangaroos/wallabies	Lateral caudal five., cephalic five., recurrent tarsal v., jugular v.	twenty–22g
Rock-wallabies	Lateral caudal v., cephalic v., recurrent tarsal 5., jugular five.	22g
Pademelons/small	wallabies Lateral caudal 5., cephalic v., recurrent tarsal v., jugular v.	22–25g
Flying-foxes	Fly 5.	25g needle/butterfly catheter
Microchiroptera	Fly five.	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 five., brachiocephalic v.	18–20g, 38 mm
Southern elephant seal	Intervertebral 5.	16g, 77 mm
Dolphin	Fluke five., dorsal fin five., pectoral fin v.	18–20g 40 mm, needle/butterfly catheter

^a: Cardiac puncture is non recommended for routine drove of blood.

v.: vein.

The vein should exist occluded to impede the period of venous blood, thus congesting the vessel and facilitating visualisation of the vein. This is achieved by awarding of a tourniquet, by digital force per unit area applied by an assistant or past using the non-preferred thumb and/or forefinger of the operator. In larger animals, having an assistant utilise digital pressure to occlude the vein is the usual technique, merely in small species, a condom 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 besides assistance in identifying the course of the vein; for larger veins, tapping the distal stop 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 judge of the needle and the size and volume of the syringe should be appropriate for the size of the vessel and the book of blood to be nerveless. These are commonly decided by the size of the brute and a guide for some species is given in Tabular array i.i. Both the needle and syringe should exist sterile. Needles should non exist reused between animals because of contagion of samples and possible manual of disease.

Align the needle, with the bevel facing upwards, and the syringe with the management of the vein and at an angle of approximately fifteen° to horizontal. Advance the needle to pierce the pare, subcutaneous tissue and vessel wall. Avoid lateral motion of the tip of the needle as laceration of the vein may result with subsequent haemorrhage and haematoma germination.

Ideally, venepuncture requires just one motility to pierce the vein, but in do some redirection of the needle may be required to place the needle inside the vein. If blood is not obtained after 1 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 motion. Another common problem is when the needle is driven besides deeply into the tissue and passes through the vein, in which instance the needle should be slowly withdrawn while creating a small-scale amount of negative force per unit area with the plunger of the syringe until claret appears in the hub of the needle.

One time the needle is aligned within the vein, draw back the plunger of the syringe using gentle, even force per unit area throughout the length of the syringe until an adequate amount of claret has been collected. Vigorous suction should be avoided every bit it may crusade haemolysis. Backlog suction may also crusade the vein to collapse, which hinders withdrawal of blood, particularly with small veins. The collector must exist patient and apply only very gentle pressure to the plunger of the syringe or utilise an alternating collection procedure. When the flow of blood from the vein is wearisome the sample may jell within the syringe, which tin be prevented by flushing the needle and syringe with an anticoagulant solution, such as heparin or ethylene-diamine-tetra-acetic acrid (EDTA), prior to apply (note that EDTA will bear on 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 exist easily visualised). In all cases, venepuncture is fabricated easier by practice.

In one case blood has been collected into the syringe, withdraw the needle and utilise digital pressure to the vein for a period of 30 seconds to 1 minute to foreclose blood flowing from the damaged vein. Remove the needle from the syringe (squirting the blood through the needle will issue in haemolysis), gently expel the blood into an appropriate container containing anticoagulant (Plate 1) and carefully mix it. Needles should non exist recapped following use, to avoid needle injuries, and both needles and syringes should exist disposed of in appropriate biohazard containers.

The blood of some species of Australasian mammals may comprise zoonotic organisms. Recently discovered viruses of flight-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 equally 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 claret tubes may be of use in larger animals, but are inappropriate for use in minor animals as the pressure of the vacuum collapses the vein and precludes withdrawal of claret.

Drove using a butterfly catheter

In some circumstances a butterfly catheter may exist more than suitable for venepuncture than a needle, peculiarly when animals are non anaesthetised and may move during the procedure. The butterfly catheter provides increased stability and in one case placed, is less likely to come up out of the vein or to lacerate it if the patient moves. There is an increased 'expressionless book' in butterfly catheters because of the tubing, making them less suitable for pocket-size animals and small veins with a deadening catamenia of claret.

Culling collection sites

Collection from arteries

Some situations may dictate that an arterial sample of blood be collected; for example, an assessment of claret gas concentration. The mechanics of collecting blood from arteries are like to those for venepuncture. After collection, apply pressure level for a longer menstruation of time following withdrawal of the needle, every bit the claret pressure of arteries is higher than in veins and significant haematoma formation may effect from leakage.

Arteries may be used for routine collection of blood samples, such every bit the cranial tibial avenue in small possums, just in some cases sampling from an artery volition be accidental. In some sites the artery may be anatomically shut to the vein, for example, the femoral artery is shut to the femoral vein, and the collector should exist suspicious that the sample of blood is arterial rather than venous when the blood is a noticeably brighter cherry-red colour and when the collection is more than rapid (considering of the college pressure of the arterial system). Information technology is also possible to puncture both vessels when the vein and artery are in close proximity, in which case a 'mixed' (arteriovenous) sample volition be collected (Tvedten et al., 2000).

Collection from the heart

Cardiac puncture may be used to collect relatively big volumes of blood from almost any species, but has potentially untoward sequelae including pulmonary haemorrhage, bleeding into the pericardial sac and cardiac tamponade, and fibrosis of the cardiac musculus. These may nowadays clinically as respiratory distress, cardiac insufficiency or sudden expiry and consequently, cardiac puncture is non 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 judge volition vary with the size of the patient. Place the anaesthetised patient on its side (lateral recumbency) and palpate the lower department of the thorax for the strongest heartbeat. When the animal'due south front end 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 about the costochondral junction and avant-garde 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 so handled as previously described. Mesothelial cells may exist inadvertently harvested from the pericardium during cardiac puncture and afterward observed in blood films (Plate ane).

Collection from peripheral veins and capillary beds

In many species a minor volume of blood may be obtained past rupturing a small peripheral vein, such every bit an ear vein or the lateral caudal vein, using a scalpel or the point of a needle. The drop of blood that wells upward into the hub of the needle may be collected into a capillary tube. Similarly, a claret sample may exist obtained from a closely clipped toenail, which is a site more usually 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 and so the distal blast 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 so 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 ii methods are not recommended as there are usually alternating methods that allow a greater volume of blood to be collected with less tissue impairment.

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 claret should not be used for comparison of the results of laboratory analysis of blood collected from capillary beds. Claret 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 every bit 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 manus, and the loose skin of the head is tightened with the thumb and center finger. With the assistance of the index finger the eye is fabricated to bulge slightly by further traction of the skin adjacent to the centre. The tip of the pipette is then placed at the lower inner corner of the centre 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 equally a useful reservoir. A slight withdrawal of the pipette frees the tip so that the accumulated blood is drawn into the tube by capillary activity. The actual bleeding role of the procedure takes about 2 seconds. Residual blood around the centre is swabbed articulate with a soft absorbent tissue to avoid clot formation. Haemorrhage unremarkably 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 claret to be collected frequently, but requires technically skilled operators, may crusade haematomas and optic nerve damage, and is becoming more than controversial (Nahas and Provost, 2002). Bradley et al. (1980) reported that repeated samples may be taken past this method with no untoward sequelae, only blindness in two Melomys spp. was believed to accept been the result of blood collection using this method (Kemper et al., 1987). This procedure is not recommended for exophthalmic animals considering ocular damage may upshot (Booth, 1999a).

Special considerations when collecting blood

Drove of blood samples in common cold climates

When blood is nerveless 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 commonly 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 exist affected by a wide range of factors, including species, torso type, body size, climate, physiological action, pregnancy and lactation (Jain, 1986), and is unremarkably reported equally millilitres per kilogram of body weight. Blood volume may exist determined by measuring both plasma book 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 decide claret book (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 greyness, common wallaroo: 87.5 mL/kg) (Maxwell et al., 1964) and the mutual 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 accept been compiled (Jain, 1986): domestic dog 77–78 mL/kg, sheep 62–66 mL/kg, equus caballus 88–110 mL/kg, rat 70–82 mL/kg and republic of guinea grunter 66–78 mL/kg. When the blood volume is not known for a item species, seventy mL/kg may be used as a reasonable guide.

The volume that may be safely nerveless (i.eastward. without challenging circulatory system homeostatic mechanisms) is dependent on the total claret book of the fauna and therefore the size (body mass) of the particular animal. Clinical signs of hypovolaemic shock occur when blood volume is decreased to 60–seventy% of 'normal' (Jain, 1986). A report of rats found no significant alteration in haematological values (haematocrit and haemoglobin and erythrocyte concentrations) when less than vii.5% of blood volumes was removed, and upward to xx% of blood volume could exist removed without adverse effects on the welfare of the beast (Nahas and Provost, 2002).

Handling of blood samples

Anticoagulants

Once the sample of claret has been collected it must exist expeditiously mixed with an anticoagalant to forestall 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 equally 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, five mL, ii mL and 0.5 mL book (Plate 2), are commercially available and the advisable sized tube should be selected for the book of blood collected. Significant underfilling of tubes may result in artefactual changes in the shape of erythrocytes (e.g. echinocytosis).

Figure one.one The 'wedge' method for making blood films. (a) A drop of blood is applied to a slide using a capillary tube. (b) A 'spreader' slide is 'reversed' into the drib of blood (which spreads laterally). (c) The spreader slide is and then moved forward and the blood flick is formed on the kickoff slide.

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 cease) the tube so that the blood is thoroughly mixed with the anticoagulant. Vigorous shaking may crusade haemolysis and should exist avoided. Accept care with small (0.five mL, 'paediatric') tubes to ensure the blood is mixed with the anticoagulant because information technology may be held stationary past 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.east. haemolysis), which may issue in spurious laboratory information such every bit decreased haematocrit and increased hateful corpuscular haemoglobin concentration. Haemolysis may besides 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).

Claret films

There are several methods that may be used to brand blood films. Prior to making any blood film the sample must be thoroughly (simply 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 demote top) then place a drop of blood (a generous 'pin-head' size) towards the finish of the slide. Concord a second slide at approximately 45 degrees to the first slide to spread the drop of blood as follows: impact the 'spreader' slide to the first slide in front of the drop of blood, opposite the 'spreader' slide into the drop of claret, pause momentarily while the blood spreads laterally towards the edges of the slide and so rapidly and smoothly propel the 'spreader' slide forrard. Some practical points to consider are:

•  if the blood drop begins to dry on the slide, it volition not spread well;

•  if the spreader slide is stopped and 'lifted' at the end of the motion picture, a thick band of claret volition grade at the 'leading edge' of the film;

•  if the blood film is besides thick or 'runs' off the edge of the slide, then too much blood has been placed on the slide;

•  if the blood film is as well 'thin' or too 'short', so 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 earlier first the forward motion of the spreader slide;

•  clots and foreign material in the sample usually appear as 'chunks' most the leading edge of the motion-picture show.

Finally, the quality of blood films produced improves with do and it is often beneficial to make several slides per sample so select the finest example for further processing.

Alternative methods to produce a blood picture include those that use 2 slides, a coverslip and a slide, or two coverslips. The 'ii 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 claret is placed on a slide (held by the operator), and then a 2nd slide (held at correct angles to the starting time slide) is flatly touched to the drop of blood (with no downward pressure level) and gently avant-garde forth the offset slide. Similarly, a encompass-slip may be used instead of the 2nd slide. Finally, when merely very pocket-sized volumes of blood are available the 'two coverslip' method is most advisable. A driblet of claret is placed on a coverslip then a second coverslip placed on top of the offset. The blood spreads under the weight of the second coverslip and when the ii are drawn apart, 2 films are produced.

The gross appearance of blood films can vary with operator and method (Plate iii

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