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Veterinary Handbook Contents

10.3 Methods For Sample Collection

10.3.1 General

The core samples, listed in Table 10.2 below, should be collected at every necropsy, and stored in buffered formalin for histology and possibly other tests. These samples will allow consistent differentiation of many of the common causes of mortality in export animals.

Table 10.2: Recommended list of core samples to be collected from every necropsy (assuming samples can be collected)

Core samples

1. Heart

2. Lung

3. Trachea

4. Liver

5. Kidney

6. Spleen

7. Ileocaecal valve

8. Ventral rumen including pillar

Other samples should be collected as directed by your laboratory diagnostician or from organs/tissues that appear abnormal in size, shape, smell, contour, colour, consistency and content during the gross necropsy.

Where possible, over collection and “banking” of collected tissues is preferred to under-collection of tissues. This is because you only get one chance at collecting the samples. Samples can undergo staged diagnostic testing at the laboratory to limit costs and conserve diagnostic resources.

Fixation of a complete set of tissues from a comprehensive necropsy of a cow, sheep or goat will usually require at least two, 2 L plastic, screw-top containers, each containing 1 L of 10 % buffered formalin solution. Lids should be retightened at intervals as they will loosen with changes in temperature.

There are difficulties in collecting, storing and processing samples from necropsies performed during an export voyage or in overseas destinations. There may be limitations on the type, or number/volume of samples that may be managed.

Importing samples back into Australia for examination will require official approval from the Department of Agriculture (DoA).

If the necropsy is being conducted as part of an animal welfare investigation, samples must be collected in triplicate. One set goes to each of the lab, owner or person-in-charge, and yourself.

Samples of organs and tissues should be collected as soon as possible after death and be promptly preserved by chilling to maximise usefulness for microbiological examination, or promptly preserved in 10 % buffered formalin for histological examination.

10.3.2 Samples For Histology

  • Place tissue in 10% buffered formalin solution at a ratio of ten parts formalin to one part tissue sample for three days at room temperature. The volume of formalin can be reduced later to facilitate transport of samples. For proper fixation, tissue samples from solid organs such as liver, kidney and spleen should not exceed 1 cm thickness.
  • Tissue samples should be collected from the margins of lesions to obtain normal and abnormal tissue in the one piece. It is good practice to also take separate pieces of normal and abnormal tissue from the same organ. Avoid cutting long, thin sections as they curl up on fixing, and are difficult to process. Blocks of 2 cm x 2 cm x 1 cm are best. Hollow viscera such as intestines should be partly incised lengthwise to allow adequate penetration of fixative to the inner surface.
  • The fibre orientation of nerves and muscle can be preserved by laying out 2 to 3 cm lengths on a flat wooden stick then fixing in formalin.

10.3.3 Samples For Microbiology

  • Samples for microbiology should be collected using sterile swabs, or instruments sterilised by boiling or autoclaving.
  • One tissue per sterile container is the rule for specimens collected for microbiological examination. Multiple tissues per container complicates the interpretation of any isolations made.
  • It is crucial not to contaminate culture containers either with formalin solution, which will kill organisms, or with materials from other carcases that could give a false positive culture.
  • A culinary gas torch can be used to heat a spatula or spoon to sear and heat sterilise surfaces of tissues. A cut can then be made into the seared area with a sterile scalpel blade and a swab inserted into the tissue below. This technique is used when uncontaminated samples of lung or liver are required for microbiology. The technique can also be applied to the surface of joints before aspirating contents with a needle and syringe including the ventral aspect of the atlanto-occipital joint to aspirate cerebrospinal fluid.

10.3.4 Samples For Serology

Blood should be collected into clot collection tubes and the tubes stood vertically at room temperature for 2-4 hours, or until the blood has separated and the clot has retracted to the base of the tube. Red cells can be discarded. The serum can then be decanted into other labelled tubes and stored chilled or frozen.

10.3.5 Samples For Toxicology

  • Collect blocks of solid organs of at least 2 cm x 2 cm x 2 cm into plastic resealable bags. Wrapping tissues in aluminium foil prevents contamination with plastic which can affect results of some toxicological analyses.
  • Collect containers or resealable bags of rumen or intestinal content of at least 30 mL and freeze.

10.3.6 Blood Smears

If blood parasites are suspected, make a blood smear on a glass slide. Cut off the tip of a dependent ear and squeeze a drop of blood on to a glass slide and make a smear. It is difficult to obtain blood from the upper ear in animals that have been bled. Try the dependent ear into which remaining blood will have pooled.

10.3.7 Multi-Test "Dipsticks"

Multi-test “dipsticks” can be used to test urine and rumen contents for pH and nitrate, and urine additionally for ketones, glucose, protein and leucocytes. Each reagent pad must be immersed by dipping the dipstick into fluid or adding fluid to the dipstick. Excess fluid should be removed to prevent dilution of the reagents or mixing of reagents between pads. This can be achieved by tilting the dipstick around its long axis causing the fluid to run off one of the long sides of the dipstick. Alternatively, place a drop of urine or rumen fluid on to the separate reagent pads on the dipstick using a syringe and needle rather than flooding the dipstick.