Copper-associated hepatitis is characterised by abnormal accumulation of copper within hepatocytes, which leads to oxidative damage and inflammation. It can be both primary (a genetic defect in copper metabolism) and secondary (due to obstruction of bile flow). Copper hepatopathy is seen most commonly in the Bedlington Terrier, Doberman Pinscher, Dalmatian, Labrador Retriever, Cocker Spaniel, West Highland White Terrier, and Skye Terrier. However, any breed can be affected.

Definitive diagnosis requires liver copper quantification interpreted in conjunction with histopathologic findings.

Specimen:

• Fresh/frozen tissue is preferred. Formalin fixed tissues can also be used but the risk of contamination is higher.
• Minimum 30mg wet weight, ideally >100mg.
• Plain, sterile, screw top polypropylene containers are preferred. Lith hep and EDTA tubes may be a source of contamination.
• Use the smallest container the tissue sample will fit in. Placing small samples in large containers can cause dehydration of the tissue and increase the apparent mineral concentration of the tissue.

Recommended container size guide:

• • Liver biopsy samples: <30 mg - place in 1.5ml container.
  • Small samples: <0.5 grams (approx. size <1cm across) – place in 1.5ml container.
  • Medium samples: 5 – 2.0 grams – place in a 5 ml container.
  • Large samples: 0 – 10.0 grams – place in a 50 ml container.

Collection protocol:

• For biopsy samples, rinse blood/blood clots from the tissue with saline. Remove excess blood/blood clots/saline with lint free wipes.
• Place the tissue into a suitable container (see above).
• Do not wrap in gauze or any other material.

Comments:

• Attempts are made to assay all samples on a wet weight basis rather than a dry weight basis. Drying samples increases the risk of contamination and adds significantly to the analysis time and cost, without contributing to the final interpretation.
• Excess blood, blood clots and saline cannot be differentiated from the original tissue. With small samples, the entire contents of the container (condensate, fluid and tissue) are used and assumed to have originated from the original tissue. Leaching from the tissue into any excess fluid and/or gauze will reduce the apparent mineral concentration of the tissue.

AMH is a hormone involved in gender differentiation in the developing embryo. It is produced by the granulosa cells of ovarian follicles and the Sertoli cells of the testicles. AMH levels markedly decline following neutering.

AMH is useful to confirm neutering status in dogs, cats and rabbits, and to identify cases of ovarian remnant syndrome and cryptorchidism.

AMH can also be used to detect granulosa cell tumours in mares and to distinguish between cryptorchid and castrated horses.

Species:

Canine, feline, laprine and equine

Specimen:

Clotted blood (1.0 mL) in plain (red top) tube

Collection protocol:

• A fasted sample is preferred to avoid lipaemia.
• Collect blood into a plain tube.
• Allow the sample to clot for 30 minutes at room temperature, then refrigerate.

Comments:

• The test is only suitable for animals over 6 months and repeat testing may be needed for animals between 6-12 months.
• Testing should be delayed for at least 30 days after spaying/neutering to allow for residual AMH concentrations to decline.
• Negative AMH test results do not rule out the possibility of ovarian remnant syndrome or residual/retained testicular tissue. If clinical signs are consistent with presence of an ovarian remnant or residual/retained testicular tissue but AMH concentrations do not support this, progesterone testing (females) or testosterone testing (males) should be considered. GnRH/hCG stimulation testing may be required in some cases.
• AMH concentration will tend to increase with sample storage therefore samples should arrive at lab within 2 days of collection.

Cushing’s syndrome is the umbrella term for a range of clinical syndromes that is caused by a chronic excess of glucocorticoid activity. Naturally occurring Cushing’s syndrome can be pituitary-dependent or adrenal-dependent. Iatrogenic causes of Cushing’s syndrome occur secondary to chronic administration of systemic or topical glucocorticoids.

The ACTH stimulation test is a screening test for hypoadrenocorticism and naturally-occurring Cushing’s syndrome and the only test that can document iatrogenic Cushing’s syndrome.

The ACTH stimulation test is considered to be less sensitive and slightly more specific than the Low Dose Dexamethasone Test (LDDST) for diagnosis of Cushing’s syndrome. It may be necessary to perform both an ACTH stimulation and a LDDST at independent time points when investigating for Cushing’s syndrome.

Resting cortisol is not a screening test for Cushing’s syndrome. Resting cortisol can be used as a screening test to rule out hypoadrenocorticism; a low value cannot prove the presence of hypoadrenocorticism and should be followed by an ACTH stimulation test for confirmation of the disease.

Species:

Canine

Specimen:

Serum (0.5 mL) or clotted blood (1.5 mL) in Gel (gold top) or plain (red top).

Collection protocol:

1. If assessing response to medical therapy, sample should be collected 4-6hrs after administration of trilostane
2. Take a baseline/0hr blood sample into a serum tube
3. Inject 5 μg/kg Synacthen® IV
4. Take another blood sample 1h later into a serum tube
5. Label sample times clearly on the tubes and submit both to the laboratory
6. If only submitting a single cortisol sample, label the time it was collected and if an ACTH stim test was performed.

Comments:

• All corticosteroids apart from dexamethasone will cross-react with the cortisol assay. To avoid a cross-reaction, the following withholding times are recommended:
  • Cortisone acetate 12h
  • Prednisolone 24h
  • Fludrocortisone 24h
• If glucocorticoid therapy is required for immediate management of a potential Addisonian dog, a single dexamethasone dose should be used as this will not interfere with the ACTH stimulation test.
• Intravenous use of the synthetic ACTH is considered standard; though intramuscular use has also been shown to be effective; for standardisation and comparability purposes IV administration is recommended, where possible
• Specific testing for hyperadrenocorticism should not be performed in unwell or significantly stressed animals, as this may lead to false positive results. In animals with concurrent disease, adrenal function testing should be delayed until current disease is controlled.
• Specific testing for hypoadrenocorticism (Addison’s disease) may be done in unwell/stressed animals.
• Anticonvulsant therapy (phenobarbital, primidone, phenytoin) may cause an elevated post-ACTH cortisol concentration.

Cushing’s syndrome is the umbrella term for a range of clinical syndromes that is caused by a chronic excess of glucocorticoid activity. Naturally occurring Cushing’s syndrome can be pituitary-dependent or adrenal-dependent. Iatrogenic causes of Cushing’s syndrome occur secondary to chronic administration of systemic or topical glucocorticoids.

The ACTH stimulation test is a screening test for hypoadrenocorticism and naturally-occurring Cushing’s syndrome and the only test that can document iatrogenic Cushing’s syndrome.

In cats, the sensitivity of the ACTH stimulation test for diagnosis of hyperadrenocorticism is poor (estimates vary from 30 – 50%). A low dose dexamethasone suppression test or the urine cortisol:creatinine ratio screening test may be preferred

Hypoadrenocorticism (Addison’s disease) is extremely rare in cats; an ACTH stimulation test should only be considered when all other potential causes for the illness being investigated have been excluded.

Species:

Feline

Specimen:

Serum (0.5 mL) or clotted blood (1.5 mL) in Gel (gold top) or plain (red top).

Collection protocol:

1. Try to maintain the cat in a stress-free environment
2. Take a baseline/0hr blood sample into a serum tube
3. Inject 125 ug/cat IM or IV
4. Collect a 30 minute post-ACTH blood sample and label it ’30 mins’.
5. Collect a 60 minute post-ACTH blood sample and label it ‘60 mins’.
6. Store samples at 4°C and submit all 3 samples together to the laboratory. If transport to the laboratory will be delayed (> 12 hours), the sample should be centrifuged and the serum separated.

Comments:

• Specific testing for hyperadrenocorticism should not be performed in unwell or significantly stressed animals. Adrenal function testing should be delayed until intercurrent disease is controlled.

• Any form of corticosteroid therapy may interfere with adrenal function tests, by (a) cross-reacting in the cortisol assay (except for dexamethasone), or (b) affecting the pituitary/adrenal axis via suppression of ACTH production. As a guide, the minimum periods for which corticosteroid therapy should be withheld before an ACTH stimulation test are:
  -Injectable (short acting) 7 days (48 h if dexamethasone, except in a potential Addisonian dog, see below).
  -Oral 2 weeks.
  -Topical 2 weeks.
  -Injectable (depot) 2 months.

Flow cytometry uses laser-based technology to sort cells based on their size, fluorescence and granularity. Certain antibodies can be applied to detect specific cell markers on round cell populations. This is most commonly used to:

- determine if a lymphocytosis is more likely to be reactive or neoplastic

- immunophenotype and potentially subtype B and T-cell lymphomas and leukaemias

- differentiate between acute and chronic leukaemias

- differentiate between lymphoid and myeloid neoplasms

Species:

• Canine
• Feline

Specimen:

• Whole blood
• Body cavity effusions
• Organ tissue aspirates

Available antibodies:

• Dogs: CD45, CD3, CD4, CD5, CD8, CD21, CD14, CD61, MHCII, MPO, CD34, cell viability marker
• Cats: CD4, CD5, CD8, CD21

Collection protocol:

Blood samples

All samples for flow cytometry must have current CBC results and clinical pathologist blood film evaluation (ideally within 2 days of flow cytometry submission).

Please submit a minimum sample volume of 1 mL blood in EDTA.

If a concurrent FBE is to be performed, please submit 2 mL of EDTA blood and two fresh blood smears (additional FBE charge will apply).

For leukaemia immunophenotyping, samples with a lymphocytosis and/or circulating atypical cell count> 5 x 10⁹ cells/L are recommended.

For lower cell counts, please discuss suitability of the sample with our clinical pathologists.

Body cavity fluid samples

All samples for flow cytometry must have a body fluid analysis and clinical pathologist evaluation within 24 hours of submission of sample for flow cytometry.

Please submit a minimum sample volume of 1 mL of fluid in EDTA and a minimum of 0.5 mL of fluid in a plain tube (no gel or SST).

If a concurrent fluid analysis is to be performed, please submit a further 1 mL of sample in EDTA and two fresh fluid smears (additional fluid cytology charges will apply).

If the fluid sample is <30g/L, please add a few drops of serum from the patient, or another animal of the same species to aid cell preservation.

Organ aspirate samples

All samples for flow cytometry must have a cytologic evaluation within 24 hours of submission of sample for flow cytometry.

Sample collection instructions:

- Place 1 mL 0.9% saline into an EDTA tube (no Serum-Z, CAT, SST or gel)

- Add 0.1 mL of serum from the patient, or another animal of the same species

- Aspirate the organ (collect from multiple areas with multiple aspiration attempts) and gently expel contents into saline/serum tube

- Rinse residual cells from the syringe by drawing up saline/serum mixture and gently expel back into the tube.

- Repeat aspiration and rinsing until the saline/serum solution looks slightly turbid

- If sample remains clear, then repeat the organ aspiration and rinsing steps

Comments:

• For optimum viability, submit samples within 48 hours of collection. Blood samples sent within 4 days and tissue samples within 72 hours from collection may be acceptable but not optimal due to reduced cell viability.
• Samples should be refrigerated immediately after collection. DO NOT FREEZE THE SAMPLES.
• Ideally, chemotherapy should be avoided in patients prior to the first immunophenotyping test when using flow cytometry, as antigens can be downregulated with treatment.
• Poorly cellular samples (including most CSF samples) may not be diagnostic.

Von Willebrand Disease (vWD) is the most common inherited haemorrhagic disorder in dogs. To date it has been recognised in more than 54 breeds of dogs, with a high prevalence of the disease (15-60 % of animals tested) in Scottish Terriers, Pembroke Welsh Corgis, Shetland Sheepdogs, Miniature Schnauzers, Rottweilers, Golden Retrievers and Dobermanns.

A deficiency in, or an abnormality of, vWf will affect primary not secondary haemostasis, and manifests as a “platelet defect”, i.e. haemorrhage from mucosal surfaces. The coagulation cascade is not usually affected, therefore the prothrombin time (PT) and activated partial thromboplastin time (APTT) are almost always normal in canine vWD.

A diagnosis of vWD and Haemophilia can potentially be made via genetic testing (see https://www.orivet.com) or via measurement of von Willebrand factor Antigen (vWf:Ag) and relevant coagulation factor concentrations.

This protocol applies to vWf:Ag and specific coagulation factor testing (e.g. Factor VIII and IX), and is performed at the University of Melbourne.

Species: Canine

Collection protocol:

• Refer to comments below for contraindications to vWf testing.
• The dog should be fasted for about 12 hours beforehand.
• A resting sample should be collected. Excitement must be minimised. Sedation can be used if needed.
• Collect a minimum 1.3mL of blood in Sodium citrate. Ensure that the tube is filled correctly to avoid sample rejection. Underfilled and overfilled tubes are unacceptable.
• Centrifuge the sample as soon as possible.
• Transfer plasma to a plain plastic tube using a plastic pipette. Label the tube with the patient name and ‘Plasma’.

**Under no circumstances should the plasma be transferred to a clot activator tube (with or without gel, labelled eg SST, CAT, serum separator, Serum-Z) or any other blood collection tube. Please do not transfer plasma back into a sodium citrate tube. A plastic urine jar is suitable if a plain plastic tube is not available. **

• The plasma should be kept cool, i.e. 4˚C, after collection and in transit to the laboratory. An esky with ice packs to keep the sample chilled is ideal. Transfer the sample to the laboratory as soon as possible (within 24 hours). If there is to be a longer delay, the sample should be frozen. Frozen plasma samples are stable at -20˚C for at least a month. Such samples must remain frozen when transferred to the laboratory, i.e. transport on dry ice.
• If a centrifuge is not available and a whole blood sample is being transported to the laboratory, the sample should be wrapped in cotton wool or newspaper and placed in an esky with ice packs. The sample should not rest directly on the ice packs as this will cause haemolysis.

Comments:

Avoid collecting blood from dogs that are in heat, pregnant or lactating. These physiological conditions may alter von Willebrand factor (vWf) values.

Avoid collecting blood from a sick animal or one that has recently had surgery or a bout of serious haemorrhage. Please wait until 6 weeks post haemorrhage to sample for vWD.   von Willebrand factor concentration may increase in response to stress, inflammation, acute haemorrhage and infection because it is an acute phase reactant.

A comprehensive information sheet for von Willebrand Disease in Dogs is available, upon request.