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.

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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.

Bile acids are produced in the liver. In dogs and cats, bile acids are released into the intestine following gall bladder contraction. Horses lack a gall bladder and therefore bile acids are continuously released from the liver into the small intestine. They are then reabsorbed into the portal blood, and removed by the liver for re-use.

Serum bile acids will be influenced by any process that affects any part of this recycling system. This may include biliary obstruction, loss of hepatic mass or function, or when portal blood is shunted away from the liver.

Post stimulation samples are generally paired with initial fasting samples, as post challenge samples may reveal impaired bile acid clearance that is not apparent with fasting samples.

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Species:

• Canine and feline
• Single bile acids can be measured in horses

Specimen:

• Serum or plasma (0.5 mL) in gel (gold top) or plain (red top) tubes
• Whole blood (1.5ml) in lithium heparin (green top) or EDTA (purple top)

Collection protocol (dogs and cats)l:

• The animal should be fasted for 12 hours.
• Collect a fasting blood sample. Clearly label the tube with ‘0 hr’.
• Feed the dog or cat a small meal to stimulate gall bladder contraction. It is recommended that pets <5kg of body weight eat at least two teaspoons of food, those that weigh more eat at least two tablespoons. Avoid overfeeding.
• Collect the post-prandial blood sample 2 hours after feeding. Clearly label the tube with ‘2 hr’.
• Transport both samples to the laboratory within 24 hours of collection.

Comments:

• Sample lipaemia and haemolysis may interfere with the results of the bile acid assay.
• Bile acid testing is NOT recommended in an animal that has hyperbilirubinaemia that is not due to haemolytic disease, since this already indicates impaired liver and/or biliary function.
• Bile acid concentrations may be falsely lower in animals younger than 16 weeks.

Cross-matching is used prior to a blood transfusion to determine if the donor’s blood is compatible with the intended recipient. In dogs and horses, naturally occurring antibody against important haemolytic red blood cells are not found. Therefore, these animals require prior sensitisation before a haemolytic reaction will occur (i.e. a previous blood transfusion). Crossmatching is not required before a first transfusion but is strongly recommended after a transfusion. Cats have naturally occurring antibody to red blood cell antigens and therefore crossmatching is essential before any transfusion.

The major cross match tests red blood cells of the donor against the serum of the recipient. It detects antibodies in the recipient against transfused red blood cell antigens from the donor. Major cross match incompatibilities may result in life-threatening transfusion reactions if that blood is transfused into the recipient.

The minor cross match detects antibodies in the donor serum to the recipient's red blood cells. Minor cross-match incompatibilities are usually not life threatening due to marked dilution of the donor antibodies following transfusion.

Species:

Canine, feline and equine

Specimen:

From the recipient and every donor:

• 2mL blood in purple-top EDTA tube
• 2mL blood in red-top plain tube

Collection protocol:

• Recipient: Collect 2 mL of blood into an EDTA tube and 2 mL of blood into a plain serum tube
• Donor/s: Collect 2 mL of blood into an EDTA tube and 2 mL of blood into a plain serum tube
• Refrigerate samples until transport to the laboratory

Comments:

• Autoagglutination can occur in animals with immune-mediated haemolytic anaemia. In these, interpretation of incompatible crossmatches is very difficult and a compatible donor may not be found.
• Take care to minimise haemolysis during sample collection, as this can invalidate results.

Bone marrow evaluation is indicated when peripheral blood abnormalities are detected, such as persistent cytopenias, or when abnormal/immature cells are seen on blood film exam. It can also be useful to stage neoplastic conditions, evaluate lytic bone lesions and investigate unexplained hypercalcaemia or a monoclonal gammopathy.

Bone marrow aspirates are best performed in addition to core biopsies. Core biopsy sections are best to evaluate marrow cellularity and assess for myelofibrosis and metastatic neoplasia, but cell morphology is best appreciated with cytology.

Collection and preparation of good quality bone marrow aspirates is a specialised procedure and referral to a specialist is recommended. Protocols for the collection of samples are available in textbooks.

Specimens:

• Multiple freshly prepared smears, as well as an EDTA-anticoagulated marrow sample is recommended.
• Submission of a concurrent peripheral blood sample is required to aid interpretation of the marrow sample.

Note that an FBE is included in the Aurora bone marrow evaluation fee.

Collection protocol:

Harvesting a bone marrow aspirate directly into a syringe containing anticoagulant will reduce the likelihood of sample coagulation and will allow time to prepare good-quality smears.

If no anticoagulant is used, smears must be prepared within seconds after bone marrow collection, because bone marrow clots rapidly.

Bone marrow aspirate collection when using an EDTA-coated syringe

1. Add 0.24 mL of sterile saline to a 4 mL EDTA tube

2. Repeat with a second 4 mL EDTA tube.

3. Using a regular needle, draw out the solution from each EDTA tube into the syringe that you will use to harvest the bone marrow into. You will likely be able to withdraw about 0.2-0.3 mL into the syringe. 

4. Attach the syringe to the bone marrow aspiration needle. Harvest the bone marrow directly into the syringe and then invert the syringe 6-8 times to ensure thorough mixing of the bone marrow sample with EDTA solution.

5. Smears should be prepared within minutes after collection to ensure optimal cell preservation.

Smear preparation

6. The bone marrow is expelled into a petri dish. Marrow particles appear as small white grains (flecks) in the blood-contaminated aspirate material.

7. The particles are collected by pipette or a haematocrit capillary tube.

8. To concentrate particles in blood-contaminated aspirates, aspirated material can be placed on one end of a glass slide, which is then held vertically. Particles tend to stick to the slide while blood runs off.

9. A second glass slide is placed across the area of particle adherence, perpendicular to the first slide. After marrow spreads between the slides, they are pulled apart in the horizontal plane.

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Comments

• Place cytology specimens in separate bags to formalin pots, as formalin fumes can render cytology slides non-diagnostic.

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

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Specimen:

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

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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.

85% of canine transitional cell (urothelial) carcinomas (TCC/UC) carry a mutation in the BRAF gene. This mutation may be detected by the CADET BRAF test in a urine sample containing as few as 10 mutant-bearing cells. 15% of canine TCC/UC lack BRAF mutation, but more than two-thirds of these have other genomic signatures detectable by a reflex test (BRAF-PLUS).

The overall sensitivity to detect canine TCC/UC is therefore > 95%.

Specificity of both BRAF and BRAF-PLUS tests is ~ 100%.

• The BRAF mutation has not been detected in non-neoplastic bladder lesions, including benign polyps and cystitis.
• The test is not impacted by the presence of red or white blood cells, protein, bacteria, or by drug therapy.

Indications

• Any dog with:
  • Haematuria, stranguria, and/or urinary incontinence with diagnostic imaging evidence of a bladder mass.
  • Recurrent, complicated, or antibiotic-resistant urinary tract infections presenting with haematuria.
• As a screening test in high-risk breeds (terriers, Shetland sheepdogs, Australian shepherds, cattle dogs, beagles, and border collies).
• During chemotherapy for TCC/UC to monitor for treatment success/relapse.

Species: Canine only

Specimen:

• Urine must be collected into a special preservative solution. To obtain a test kit please call the laboratory and request a CADET® BRAF urine collection kit. The test kit consists of:
  • Urine sample jar containing preservative
  • Antech instruction sheet regarding collection
  • Antech checklist form for veterinarian to complete
• The required sample is 40 mL free-catch (voided) urine
  • A smaller volume of urine (10-25 mL) may be submitted though the sensitivity may be decreased (this is less likely to be a problem if a bladder mass can be visualised).
  • A smaller volume may also limit the option of the second level BRAF-PLUS test, if required.
  • Urine collected by cystocentesis or catheterisation is NOT recommended since the sensitivity may be decreased.

Collection protocol:

• Free catch urine should first be collected into a clean, dry container and transferred into the preservative container within 15 minutes of collection.
• Once in the preservative, urine is stable for several days at room temperature when kept out of direct sunlight.
• Short periods of refrigeration should not affect the specimen but refrigeration is not necessary.
• Urine collection may take place over 2-3 days as long as each aliquot is promptly placed into the preservative solution and stored out of direct sunlight.

Comments:

• If the submitted sample does not have detectable BRAF mutation, the BRAF-PLUS test will automatically be performed at no additional charge. However, because more DNA is required to perform the BRAF-PLUS assay, a small number of samples that are BRAF-undetected will not be eligible for the BRAF-PLUS assay, and a new urine sample must be submitted (which will incur a second charge).
• When a mass is detected, histologic or cytologic confirmation of TCC/UC is recommended. Further imaging and evaluation of local lymph nodes will help to stage the disease.

Measurement of blood concentrations of cyclosporine, an immunosuppressant drug used for the treatment immune-mediated disease. 

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Species:

Canine, feline.

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Specimen:

Minimum 1mL of whole blood in EDTA (purple top) tube

Collection protocol:

• Collect blood into an EDTA tube and keep refrigerated until collection.
• Trough concentration
  • Trough samples should be collected approximately 1 hour before the next scheduled dose. 
• Peak concentration
  • Peak samples should be collected 2 hours after administration. 

Comments:

• Cyclosporin concentrations are typically checked 1-2 days following initiation of therapy, and then every 2-4 weeks.
• Monitoring of cyclosporine trough concentrations is more commonly performed in veterinary medicine.

Trough concentrations

- Pharmacodynamic studies have shown that attainment of a trough blood cyclosporin concentration of 600 ug/L reliably causes substantial immune system suppression. 

- For atopic dermatitis, response to cyclosporin treatment does not appear to be related to blood drug concentrations.

- For chronic inflammatory diseases, such as inflammatory bowel disease, a trough concentration of 250 ug/L is advised.

- For anal furunculosis, a trough concentration of 100 - 600 ug/L has been recommended.

Peak concentrations

• Peak concentrations are often 2- to 8-fold higher than trough concentrations in normal animals.
• Typically, peak blood concentrations are between 600 and 1,200 ug/L after a standard 5 mg/kg microemulsified oral dose in dogs. Peak concentrations have not been published in cats.

Equine: To aid in the diagnosis of pituitary pars intermedia dysfunction.

Canine: To differentiate between pituitary-dependent hyperadrenocorticism and adrenal-dependent hyperadrenocorticism in dogs with confirmed hyperadrenocorticism. ACTH concentrations are low when adrenal tumours are present, and normal-to-increased with pituitary-dependent disease.

Note: ACTH is a very labile protein. Suboptimal specimen collection and handling may result in a falsely low measured ACTH concentration due to exposure to red blood cells, heat and glass.

ACTH in chilled EDTA whole blood is reported to be stable for 36hrs in horses, and 90 minutes in dogs. Frozen plasma samples are preferred as they are stable for many weeks.

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Specimen:

• 5mL of frozen plasma in plain plastic tube (preferred)

Or

• 1mL of chilled whole blood in EDTA

Collection protocol:

If centrifuge available:

• Collect blood into an EDTA tube, ensuring that the tube is filled to the line and gently mix.
• Centrifuge the sample immediately.
• Transfer the plasma into a plain plastic tube (no additive) and freeze immediately.
• Samples should ideally be sent frozen with ice packs, and should still be frozen upon arrival at the laboratory.
• Note the collection date and time on the submission form.

If no centrifuge available:

• Collect blood into an EDTA tube, ensuring that the tube is filled to the line. Gently mix by inversion.
• EDTA whole blood should be refrigerated immediately and sent to arrive at the lab as soon as possible. DO NOT freeze the unseparated EDTA sample.
• Note the collection date and time on the submission form.

Comments:

• Ideally, dogs should be fasted overnight prior to collection.
• The patient should not be stressed or excited.
• Markedly lipaemic and/or haemolysed samples may yield false results and samples should be redrawn prior to submission.

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).

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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.

Fructosamine concentrations provide a reflection of blood glucose concentrations during the previous 1-3 weeks. Therefore, fructosamine is used for the diagnosis and monitoring of diabetes mellitus in both cats and dogs. In cats, fructosamine is particularly useful to differentiate between a stress hyperglycaemia and diabetes mellitus.

Serum fructosamine may also be useful for demonstrating prolonged hypoglycaemia in animals presenting with a suspicion of an insulinoma.

Species:

Canine and feline.

Specimen:

Minimum 1.5mL of clotted blood (or 0.5mL of serum) in plain (red-top) or gel (gold-top) tube

1.5mL of whole blood in lith hep (green-top) or EDTA (purple-top) can also be used.

Collection protocol:

• It is preferable to fast the animal for 12 hours prior to sample collection, though a random sample can be used if necessary.
• Collect blood sample.
• Store sample at 4°C. If transport to the laboratory will be delayed (> 12 hours), the sample should be centrifuged and the serum separated.

Comments:

• Some conditions can affect fructosamine concentrations, including hypoalbuminaemia and thyroid disorders.
• Significant decreases in fructosamine concentration can occur if samples are kept at room temperature for more than a few hours.

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

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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.

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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.

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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

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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.

Ionised calcium (iCa²⁺) is the metabolically active form of calcium and this is the form that the body senses and responds to. Approximately 50% of serum calcium is in the free ionised form.

Measurement of iCa²⁺ is a more accurate reflection of the physiological calcium state than total calcium, as it’s not affected by serum protein concentration.

Species:

Any

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Specimen:

Fill tube as much as possible.

Clotted blood in SST (serum separation tubes that contains gel) vacutainer tube.

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Collection protocol:

• The whole blood sample should be placed into vacutainer tube without uncapping lid (i.e. by inserting the needle through the cap of the tube).
• The sample should then be gently inverted 8-10 times and centrifuged 15-30 minutes after collection.
• Make sure that tube is not uncapped at any stage prior to testing.
• Wrap the SST sample (after having centrifuged the sample) in “glad-wrap” or tape the cap shut, place in specimen collection bag and keep refrigerated. Freezing is not required.

Comments

• iCa concentrations are affected by pH. Storage at room temperature or refrigeration usually results in falling pH over time which increases iCa values. Freezing and exposure to air results in increased pH and reduced ionised calcium values. Therefore, avoid uncapping sample at any point.
• iCa in whole blood (i.e. not separated) is stable for 3 hours when kept refrigerated. It is not stable in whole blood when kept at room temperature.
• If additional testing is required for the patient, please submit additional samples in a separate submission bag with a separate request form.

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.

This test is used to diagnose hyperadrenocorticism (Cushing’s syndrome) in dogs by assessing the lack of suppression of the hypothalamic-pituitary-adrenal axis in response to exogenous corticosteroids. If results of pituitary-adrenal testing confirm hyperadrenocorticism, the LDDST results can also allow differentiation of pituitary-dependent from adrenal-dependent disease.

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Species:

Canine

For feline protocol, see comments.

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Specimens:

Serum (0.5 mL) or clotted blood (1.5 mL) in gel (gold-top) or plain (red-top) tubes.

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Collection protocol:

• Collect a resting blood sample and label it with patient details and ‘0 hr’.
• Administer dexamethasone sodium phosphate at 0.01 mg/kg IV.
• Collect a 4-hour post-dexamethasone blood sample and label it with patient details and ‘4 hr’.
• Collect an 8-hour post-dexamethasone blood sample and label it ‘8 hr’.
• Stores samples at 4°C and submit all three samples together to the laboratory. If transport to the laboratory will be delayed (> 12 hours), the samples should be centrifuged and the serum separated.

Comments

• • Patients should be maintained in a stress-free environment for the duration of the test. Avoid testing in sick or stressed animals.
  • For cats, the same protocol applies, except the dexamethasone dose is 10x higher (0.1mg/kg IV).
  • 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.

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.

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Species: Canine

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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.

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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.

Measurement of serum concentrations of potassium bromide, most commonly used in the treatment for seizures.

Specimen:

Minimum 1.5mL of clotted blood (or 0.5mL of serum) in plain red top tube.

Note: Gel/serum separator tubes are NOT appropriate for this test.

Collection protocol:

• Fast the patient for 12 hours.
• Collect sample within one hour of the next scheduled dose. Once steady-state has been achieved, samples can be collected at any time point >2 hours after dosing.
• Store sample at 4°C. If transport to the laboratory will be delayed (> 12 hours), the sample should be centrifuged and the serum separated.

Comments:

• Gel/serum separator tubes are not recommended as the gel may absorb some of the drug, artificially lowering the serum concentration.
• Steady-state is usually achieved after 3-5 months of therapy.
• Recommendations for when to assess potassium bromide concentration:
  • 1 and 3 months after commencement of therapy
  • 1 month after a change in dose
  • Every 12 months for long-term therapy
  • When signs of dose-related toxicity develop, such as ataxia or sedation
  • If > 3 seizures occur between these times

Measurement of serum concentrations of phenobarbitone, most commonly used in the treatment for seizures.

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Species:

Canine, feline.

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Specimen:

Minimum 1.5mL of clotted blood (or 0.5mL of serum) in plain red top tube.

Note: Gel/SST are NOT appropriate for this test.

Collection protocol:

• Fast the patient for 12 hours.
• Collect blood and place into serum tube.
• Store sample at 4°C. If transport to the laboratory will be delayed (> 12 hours), the sample should be centrifuged and the serum separated.

Comments:

• Gel/serum separator tubes are not recommended as the gel may absorb some of the drug, artificially lowering the serum concentration.
• Steady-state serum and tissue phenobarbital concentrations are achieved after 7-10 days of therapy.
• There is no therapeutically relevant change in serum phenobarbital concentrations throughout a daily dosing interval in most epileptic dogs. Therefore, timing is not important when collecting blood samples to measure serum phenobarbital concentrations in most epileptic dogs treated long-term with phenobarbital. However, trough blood samples may be beneficial to maintain consistency in interpretation, and are best taken in the early morning, before dosing, in a fasted dog.
• Induction of liver enzymes (ALP and to a lesser extent ALT) is common in animals receiving phenobarbital therapy.
• Side-effects of phenobarbital administration may include hepatotoxicity and blood dyscrasias. In addition to monitoring serum phenobarbital concentration, assessment of a CBC and biochemistry panel is recommended every 6-12 months. If there is concern for liver dysfunction, assessment of fasting and post-prandial bile acids is also recommended.

Recommendations for when to assess phenobarbital concentration:

• 2 and 6 weeks after commencement of therapy
• 2 weeks after any change in dose
• If significant side effects develop
• Every 6 months when seizures are well controlled
• If >2 seizure events occur between these times

The urine cortisol/creatinine ratio (UCCR) is non-invasive and sensitive test to exclude Cushing’s disease in dogs. However, it is not very specific, as high UCCR results can occur with hyperadrenocorticism, stress or non-adrenal illness.

It can also be useful to exclude hypoadrenocorticism in dogs.

It is not a useful test to monitor the response to therapy in dogs with hyperadrenocorticism.

.

Species:

Canine

Specimens:

2mL of urine in a urine pot.

Collection protocol:

• Attempt to maintain the dog in a stress-free environment for 24 h prior to urine collection.
• Collect a free-catch urine sample, preferably in the morning.
• Refrigerate and submit to laboratory.

Comments

• Screening for hyperadrenocorticism should not be performed in unwell or significantly stressed animals.
• Corticosteroid-containing medications may interfere with adrenal function tests, including the UCCR.