Insulin dysregulation occurs when there are abnormalities in insulin metabolism leading to resting hyperinsulinemia, postprandial hyperinsulinemia, or tissue insulin resistance. Hyperinsulinemia may develop due to increased insulin secretion or decreased insulin clearance. Detection of hyperinsulinemia is important as it is associated with an increased risk of laminitis. For practical clinical application, detection of insulin dysregulation may be made by measurement of insulin in a baseline (resting) serum sample; however, dynamic testing may be necessary to detect insulin dysregulation in some horses. Insulin dysregulation is a central component of equine metabolic syndrome (EMS) but may also exist in association with pituitary pars intermedia dysfunction (PPID), systemic illness, pregnancy, stress, pain, and starvation.

Equine metabolic syndrome (EMS) is an endocrine condition characterized by generalized or regional (cresty neck) adiposity, insulin dysregulation, and either clinical or subclinical laminitis. Horses with EMS tend to be middle aged (< 15 years). The physical appearance is one of generalized or regional obesity. These horses are at increased risk for development of laminitis due to the increased adiposity. Contributing factors include diets of a high glycemic index (lush new pasture or grain diet), limited exercise, and genetic predisposition. Diagnosis and management of EMS before the onset of laminitis is important for a successful outcome.

• Do not feed grain or other rapidly absorbed carbohydrate within four hours of collection. Horses do not need to be fasted. A serum sample may be collected following consumption of hay or pasture.
• False increases in insulin concentration may be seen during an acute onset of a painful condition (such as laminitis). It is recommended that measurement of insulin be postponed until the pain has subsided.
• Collect a resting serum sample for measurement of insulin and glucose concentrations.
• Keep sample chilled. Separate serum from cells within four hours of collection as a false decrease in glucose concentration is expected when the sample is not separated promptly. Store serum either refrigerated (no longer than overnight) or frozen. Ship sample overnight with ice packs.

• The oral sugar test (OST) provides a more sensitive assessment of insulin and glucose metabolism compared to measuring resting insulin and glucose concentrations. This test can be performed as an initial screening test or when a baseline (resting) serum sample is inconclusive but insulin dysregulation continues to be suspected.
• The night prior to testing leave only 1 flake of hay in the stall or dirt pen before midnight.
• The following morning collect a baseline serum sample for measurement of insulin and glucose concentrations.
• Administer 0.15 mL/kg Karo® Light Syrup orally using 60-mL catheter-tip syringes (e.g. 75 mL per 500 kg).
• Administration of 0.45 mL/kg Karo® Light Syrup orally using 60-mL catheter tip syringes (e.g. 225 mL per 500 kg) may improve test performance in horses with normal response to the lower dose of Karo® Light Syrup but still suspected to have insulin dysregulation.
• Collect a serum sample at 60 and/or 90 minutes after Karo® Light Syrup administration for measurement of insulin concentration.
• Keep samples chilled. Separate serum from cells within four hours of collection as a false decrease in glucose concentration is expected when the sample is not separated promptly. Store serum either refrigerated (no longer than overnight) or frozen. Ship samples overnight with ice packs.

Pituitary pars intermedia dysfunction (PPID), formerly referred to as Equine Cushing’s Syndrome, occurs when oxidative stress reduces tonic dopaminergic inhibition of pars intermedia melanotropes, giving rise to hyperplasia, microadenomas, and macroadenomas. At the time of diagnosis, horses with PPID are typically > 15 years of age, and usually older than 20 years of age. Clinical signs vary depending on the stage of disease. Common clinical signs include hypertrichosis, delayed shedding, loss of topline musculature, abnormal sweating, and chronic laminitis.

Detecting excessive endogenous plasma adrenocorticotropic hormone (ACTH) derived from the abnormal pars intermedia is the most common diagnostic test for PPID. The sensitivity of endogenous ACTH is acceptably high in equids with moderate to advanced PPID; however, in early cases this test may yield a false negative result. For early cases of PPID, a more sensitive option is the thyrotropin-releasing hormone (TRH) stimulation test (measuring ACTH response). The overnight dexamethasone suppression test (ODST) may still be performed; however, this test is falling out of favor due to its laboriousness and lower sensitivity compared to measurement of endogenous ACTH. Regardless of the diagnostic test chosen, it is important to interpret all laboratory test results in conjunction with the horse’s signalment, history, and clinical signs. In addition, testing for insulin dysregulation is recommended in all PPID cases.

• Endogenous ACTH has supplanted endogenous cortisol measurement (either resting or dynamic testing).
• Endogenous ACTH concentrations may be falsely increased due to severe systemic illness, excitement, stress, exercise, or time of year.
• It is recommended that medication, such as pergolide, be given as normal when monitoring the effectiveness of treatment. A lower endogenous ACTH concentration compared with pre-treatment results is suggestive of a favorable biochemical response to treatment; however, it is recommended that dose adjustments be based on clinical response in conjunction with biochemical measurements.

• Collect EDTA plasma at any time of day.
• Keep sample chilled. Separate plasma from cells within four hours of collection. Store plasma either refrigerated (no longer than overnight) or frozen. Ship sample overnight with ice packs. Factors that most affect ACTH degradation and a false decrease in the measured concentration include heat and time spent on the red cells.

• Do not perform within 12 hours after an oral sugar test (OST) or a grain meal due to blunting of pituitary ACTH responses to TRH.
• Collect a baseline EDTA plasma sample for ACTH measurement.
• Administer 0.5 mg (equids < 250 kg) or 1.0 mg (equids > 250 kg) of TRH intravenously (IV).
• Collect an EDTA plasma sample for ACTH measurement exactly 10 minutes after TRH administration.
• Keep sample chilled. Separate plasma from cells within four hours of collection. Store plasma either refrigerated (no longer than overnight) or frozen. Ship samples overnight with ice packs. Factors that most affect ACTH concentrations include heat and time spent on the red cells. Proper handling and shipping of the sample is critical to minimize ACTH degradation and prevent a false decrease in the result.
• From July to December, this test should only be used to identify negative cases. Many false positives are noted due to the influence of season.

• Collect a baseline serum sample in the late afternoon for cortisol measurement.
• Administer dexamethasone at 40 µg/kg IM (20 mg for a 500 kg horse)
• Collect a post-dexamethasone serum sample at 15 hours and 19 hours for cortisol measurement. Alternatively, a single post-dexamethasone serum sample may be collected between 15 – 19 hours for cortisol measurement.
• Keep sample chilled. Separate serum from cells within four hours of collection. Store serum either refrigerated (no longer than overnight) or frozen. Ship samples overnight with ice packs.