Glucose is derived from digestion of dietary carbohydrates, breakdown of glycogen in the liver (glycogenolysis, this provides stores for maintaining glucose in blood during fasting or food-deprived states – until stores get depleted, of course, then gluconeogenesis takes over) and production of glucose from amino acid precursors in the liver (gluconeogenesis). Note that the kidney can also be a source of gluconeogenesis. In ruminants, the main source of glucose is gluconeogenesis from volatile fatty acids (propionate) absorbed from rumen by bacterial fermentation. Glucose is the principal source of energy for mammalian cells. Uptake in most cells is mediated by a group of membrane transport proteins, called glucose transporters (GLU), some of which are insulin-dependent (particularly muscle and fat), e.g. GLU-4. There are glucose receptors in the the liver, red blood cells and brain that do not require insulin, preserving metabolic function of these vital organs.


Blood glucose concentration is influenced by hormones which facilitate its entry into or removal from the circulation. The hormones affect glucose concentrations by modifying glucose uptake by cells (for energy production), promoting or inhibiting gluconeogenesis, or affecting glycogenesis (glycogen production) and glycogenolysis and are listed below. The most important hormone involved in glucose metabolism is insulin, which enables energy use and storage and decreases blood glucose concentration. Several hormones oppose the action of insulin and, therefore, will increase blood glucose. The main hormones that mediate this effect are glucagon (fasting states), growth hormone, catecholamines, and corticosteroids. The increase in blood glucose can occur through inhibition of insulin release, stimulation of glucose-yielding pathways (glycogenolysis, gluconeogenesis), or decrease of glucose uptake or use by tissues. Table 1 below summarizes these effects. Collectively, increases in these insulin opposing hormones can induce a state of insulin resistance. Insulin resistance can also be mediated by inflammatory cytokines (tumor necrosis factor-α [TNF-α]), obesity and pregnancy (Weiser et al 2013). Inflammatory cytokines are thought to be responsible for insulin resistance observed in sepsis. Hyperglycemia in critical care patients has been associated with a poor outcome and has prompted the use of glucose monitoring in such patients in human and veterinary medicine. In pregnancy, hormones such as progesterone can cause insulin resistance (this is thought to be mediated through growth hormone release) and results in gestational diabetes in humans. Pregnancy-associated hormones may also contribute to insulin resistance and hyperlipidemic syndromes in pregnant horses, ponies and camelids.

  • Insulin: Insulin is produced by β cells in the pancreatic islets. Insulin release is stimulated by glucose and amino acids, among other compounds, including fatty acids, glucagon, and incretins released by the gastrointestinal system, such as gastric inhibitory polypeptide and glucagon-like peptide 1. Glucose is the main stimulus, however, and works via being taken up gy GLU-2 receptors in islet cells, which then stimulates insulin release (performed and newly produced) via depolarizing the cell membrane, causing intracellular uptake of calcium and granule release. Release is inhibited by hypoglycemia, somatostatin, fasting, exercise, and alpha-adrenergic agonists, such as norepinephrine. Insulin decreases blood glucose by promoting glucose uptake through GLU-4, and its use in metabolism (e.g. energy production, protein production) by muscle and other tissue cells, such as adipose tissue. Insulin also inhibits glucose production by inhibiting gluconeogenesis and glycogenolysis. Insulin increases fatty acid and triglyceride synthesis (through stimulation of  lipoprotein lipase on endothelial cells), thus increasing fat stores (adipogenesis), and enhances glycogen synthesis and storage in the liver. Insulin induces the cellular uptake of K+, phosphate and Mg2+. Thus, insulin is pro-fat storage and pro-glucose usage. Counter regulatory hormones and drugs can antagonize the action of insulin and/or enhance glucose generation (eg corticosteroids, glucagon etc). Insulin also induces satiety, decreasing food intake through direct effects on the hypothalamus.
  • Glucagon: Glucagon causes an increase in blood glucose, by stimulating gluconeogenesis and glycogenolysis and facilitating glucose release from hepatocytes. Low blood glucose is the main stimulus for glucagon release from α cells in pancreatic islets. The glucagon release helps maintain blood glucose under fasting or food-deprived states.
  • Catecholamines (epinephrine/norepinephrine): Epinephrine from the adrenal medulla acts via β2-adrenergic receptors, whereas norepinephrine is released from nerve endings and acts on α2-adrenergic receptors. Norepinephrine and epinephrine have somewhat opposing effects on insulin release (norepinephrine inhibits, epinephrine stimulates), but the net effect of both is increased blood glucose. This occurs via stimulation of glycogenolysis (skeletal muscle; facilitated by glucose) and release of glucose from hepatocytes (epinephrine), and indirectly through inhibition of insulin release (norepinephrine), and release of growth hormone (epinephrine) and adrenocorticotropic hormone (ACTH; which increases cortisol). Epinephrine also stimulates glucagon release. The increase in glucose in response to catecholamines is usually transient (primarily due to intermittent release of catecholamines) but can be quite pronounced in cats (e.g. around 300-400 mg/dL), cattle (may be glucocorticoid versus epinephrine-mediated) and camelids (presumably epinephrine mediated). Generally, dogs develop less severe hyperglycemia (usually < 300 mg/dL).
  • Growth hormone (GH): This increases blood glucose by inhibiting glucose uptake by cells (muscle and fat). It also promotes glycogenolysis in muscle tissue. Progesterone (during pregnancy and just after parturition) may cause insulin resistance by stimulating secretion of GH. Growth hormone is released from the pituitary by growth hormone-releasing hormone, which is secreted by the hypothalamus usually in response to low blood glucose and epinephrine.
  • Corticosteroids: These increase blood glucose by inducing glucose release from hepatocytes and inhibiting glucose uptake by cells (through decreasing GLU-4). Glucocorticoids also stimulate gluconeogenesis and glucagon secretion (which also increases blood glucose). They also stimulate glycogen synthesis, resulting in the characteristic vacuolar change in the liver of dogs (wispy rarefaction), which is due to excess glycogen. Corticosteroids can also indirectly increase glucose concentrations, by providing more substrate (amino acids, certain fatty acids) for gluconeogenesis through their catabolic effects on protein and lipolytic effect on fat stores. In states of chronic stress, stimulation of hepatic gluconeogenesis by excess glucocorticoids can cause high concentrations of glucose in sick cattle (around 300-500 mg/dL but as high as 1000 mg/dL) (e.g. proximal duodenal obstruction; see below).
Hormones that influence glucose concentration
Hormone Glycogen
Glucose uptake
Effect on glucose concentration
Insulin Synthesis Stimulates
Glucagon Breakdown No effect
Catecholamines Breakdown
(indirect by
insulin inhibition)

(indirect through GH/insulin inhibition)
Transient ↑
Growth hormone Breakdown  
Corticosteroids Synthesis


Method of measurement

Several methods are currently available for measuring glucose concentration in body fluids. The most commonly used techniques are enzymatic and highly specific for glucose or its derivative glucose-6-phosphate. Glucose levels are generally determined by using the hexokinase, glucose oxidase, or glucose dehydrogenase assays. The hexokinase method is considered to be the standard for quantifying glucose concentration in body fluids and is the method used at Cornell University.

Reaction Type

Blanked end-point reaction (2-point)

Procedure for hexokinase method

In this two stage reaction, hexokinase (HK) first catalyzes the phosphorylation of glucose in the sample by adenine triphosphate (ATP) forming glucose-6-phosphate in the presence of Mg2+. In the second part of the reaction, glucose-6-phosphate dehydrogenase (G6PD) oxidizes glucose-6-phosphate to 6-phosphogluconate, in the presence of nicotinamide adenine dinucleotide phosphate (NADP+). The quantity of NADPH produced in the second reaction is measured photometrically by absorbance at 340 nm and directly correlates to the amount of glucose in the sample. These two reactions are shown below:

Glucose+ ATP hexokinase, Mg2+ > Glucose-6-phosphate + ADP

Glucose-6-phosphate + NADP+    G6PD   6-phosphogluconate +NADPH + H+

Units of Measurement

Blood glucose concentration in body fluids is measured in mg/dL (conventional units) or mmol/L  (SI unit). The conversion formula is shown below:

mg/dL x 0.0555 = mmol/L

Sample Considerations

Sample Type

Plasma, serum, urine, and body cavity fluids (including cerebrospinal fluid)

Suitable anticoagulants

Heparin, EDTA or citrate.

We do not recommend the use of fluoride oxalate tubes. Although sodium fluoride (NaF) inhibits glycolysis and stabilizes glucose levels in blood samples, sodium fluoride is hypertonic and causes lysis of red blood cells (RBC). This releases intra-RBC water which dilutes the glucose concentration. Glucose concentrations in fluoride oxalate samples are consequently lower than in promptly separated serum samples (by approximately 7-12%).

Stability with storage

  • Plasma/Serum: At 25°C, if plasma or serum is in contact with the cellular constituents of blood prior to centrifugation, the glucose concentration will decrease due to glycolysis by cellular constituents (note that these changes are NOT prevented by the use of serum separator tubes, i.e. the silicon does not always prevent cells from consuming glucose, unless the serum is removed from cells). Glucose losses are accelerated in blood samples collected from patients under the conditions listed below. The effects from these variables may be minimized by collecting blood into sterile vacutainer-brand tubes, promptly separating serum or plasma from cells and storing samples in a cool environment (4°C). A study with horse, dog and alpaca blood showed that glucose was not significantly decreased in blood collected into non-anticoagulant glass tubes and stored for up to 8 hours at 4°C in the horse and camelid (n=28 and 20, respectively), whereas glucose was only stable in whole blood collected into non-anticoagulant plastic tubes for 4 hours and had significantly decreased by 8% (maximum mean decrease of 12%) by 8 hours in dogs (n=26) (Collicutt et al 2015).
    • Marked leukocytosis, erythrocytosis, or thrombocytosis: These cells consume glucose in the tube.
    • Bacterial contamination of the sample: Bacteria consume glucose.
    • Storage temperature: Glycolysis is enhanced at higher temperatures. The above referenced study showed that glucose decreased by 2 hours in whole blood at 25°C by 4, 11 and 25%, by 8, 23 and 49% and by 2, 8 and 26% in horses, dogs and camelids after 2, 4, and 8 hours of storage, respectively.
  • Urine: Specimen must be stored at 4°C to circumvent room temperature-associated losses of glucose, which may reach as high as 40% after a 24-hour time period in human samples. Similar information is not available for animals.
  • Body cavity fluids: These may be contaminated with bacteria and usually contains other cellular constituents (leukocytes, RBC) which consume glucose. For accurate glucose concentrations, these samples should be analyzed immediately. If a delay in analysis is anticipated, a portion of the sample should be removed and separated from cells, as would be done for blood samples (the remaining un-spun sample can be used for cytologic analysis etc). This will however, not remove bacteria, which can still consume glucose.

Recommendations for sample submission for glucose measurement

  • Collect blood into tubes containing no anticoagulant (red top), particularly if chemistry tests other than glucose are required. Heparin anticoagulant is recommended for plasma samples.
  • Separate plasma or serum from cells promptly. This includes serum separator tubes.
  • Keep serum or plasma cool until sample submission.


  • Lipemia, hemolysis, icterus: These have minimal effect on glucose results.

Test interpretation


Sustained hyperglycemia has pathologic consequences, by causing glycosylation of protein groups. The first change is the nonenzymatic addition of glucose to protein amino groups to form Amadori products. These reach a steady state over time and do not accumulate further. Amadori products are formed with albumin (glycosylated albumin can be measured in the fructosamine assay), hemoglobin (resulting in the glycosylated hemoglobin assay, which is hardly ever used anymore) and lipoproteins (low density lipoproteins). This is a reversible change that requires several days of sustained hyperglycemia (the length of time required to form these glycosylated products depends on the degree of hyperglycemia, e.g. it takes 3 days for fructosamine to increase in cats with hyperglycemia > 400 mg/dL). 

With time, the Amadori products get transformed by dehydration, condensation, fragmentation, oxidation and cyclization to advanced glycosylation end products (AGE). This is an irreversible change. AGE form on proteins, lipids and nucleic acids and are thought to be responsible for the side-effects associated with diabetes mellitus, including diabetic neuropathy, retinopathy and nephropathy. Therefore, it is important to maintain glucose concentrations within reference intervals when treating diabetic patients.

Any cause of hyperglycemia (transient or sustained) may result in glucosuria if glucose concentrations are high enough to exceed the renal threshold. The renal threshold for glucose is species-dependent and is reported to be the following:

  • Dogs: 180-200 mg/dL
  • Cats: 280-290 mg/dL in cats (lower thresholds may occur in diabetic cats, around 200 mg/dL). Some, but not all cats, with a stress hyperglycemia may have glucosuria.
  • Horses: 160-180 mg/dL
  • Cattle: 100-140 mg/dL.

The renal threshold for camelids is unknown, but likely similar to horses and cattle.

Causes of hyperglycemia: The most common cause of an isolated hyperglycemia is “stress” (physiologic).

  • Physiologic:
    • Post-prandial: A mild physiologic hyperglycemia occurs post-prandially (this should normalize within a few hours), although this increase was only seen in 17 dogs between 2-8 hours after eating and only 6 dogs had values above the reference interval. The median glucose concentration appeared to peak at 4 hours in the study (Yi et al 2022). In liver disease, a prolonged postprandial hyperglycemia may be observed.
    • Stress“: A hyperglycemia also occurs in response to stress in all species. This can be mediated by epinephrine (and is transient, lasting 4-6 hours) or corticosteroids (results in a more sustained increase in glucose, which can last several days as long as the corticosteroids are still increased in blood). In goats, intravenous epinephrine injection (2 mg) caused an increase in mean glucose from around 50 mg/dL to around 140 mg/dL at 15 minutes. The glucose concentration was still increased 2 hours after injection (Abdelatif and Abdalla 2012). Cats and cattle tend to produce marked stress hyperglycemias. In cattle, a very high glucose (> 500 mg/dL) is a poor prognostic indicator. Note, although the single term “stress” is used to indicate a transient hyperglycemia, this can occur secondary to epinephrine or endogenous corticosteroid release (whereas a “stress” leukogram specifically refers to endogenous corticosteroid release only). Both epinephrine and endogenous corticosteroids could be contributing to a hyperglycemia in a sick animal. A stress hyperglycemia can potentially result in a glucosuria if the glucose in blood exceeds the renal threshold (thus, glucosuria alone in a single urine sample does not necessarily confirm diabetes mellitus). Internal studies at Cornell University show that cats can develop glucosuria secondary to stress hyperglycemia, however this is expected to be transient. A consistent glucosuria would be unexpected with a stress hyperglycemia and would be more compatible with diabetes mellitus in an animal with hyperglycemia that consistently exceeds the renal threshold or renal tubular damage (preventing glucose absorption from the urine filtrate).
    • Pregnancy: Late pregnancy can result in insulin resistance due to progesterone stimulating growth hormone release. The hyperglycemia usually resolves after parturition but may persist in some animals.
    • Iatrogenic: Various drugs such as xylazine, detomidine, propanalol, megestrol acetate, and ketamine can induce hyperglycemia through inducing a state of insulin resistance (frequently via release of counter-regulatory hormones), inhibiting insulin release or stimulating glycolysis or gluconeogenesis. Intravenous glucose administration can also cause hyperglycemia. Corticosteroids can also increase glucose concentrations, but this varies between studies. Mean serum glucose concentrations did not increase in 8 Beagle dogs treated for 5 days with doses of 0.5, 1, and 2 mg/kg prednisone, but glucose concentrations increased by a mean of 18±20% in dogs treated with 4.4 mg/kg/d prednisone (Tinklenburg et al 2020). Immunosuppressive doses of prednisone (4.4 mg/kg/d) or dexamethasone (0.55 mg/kg/d) to 7 cats approximately doubled the mean glucose concentrations after 56 days, but an untreated control group of cats was lacking (Lowe et al 2008). In contrast, 10 allergic cats given 1-2 mg/kg of prednisone for 13 days did not have increases in mean serum glucose concentrations (Khelik et al 2019).
  • Pathophysiologic: Sustained increases in glucose can be seen with insulin deficiency (type I diabetes mellitus) or insulin resistance (type II diabetes mellitus). Insulin resistance can be a result of increased concentrations of counter-regulatory hormones (e.g. glucocorticoids, growth hormone, progesterone) or inflammatory cytokines (TNF-α) that oppose insulin release or the action of insulin on peripheral tissues. Obesity is also associated with insulin resistance, particularly in cats and horses (as part of equine metabolic syndrome), although animals are not usually hyperglycemic as a consequence of this. Adipose tissue is now known to be an endocrine organ and can produce specific hormones (e.g. leptin, adiponectin) and inflammatory cytokines (TNF-α).
    • Sustained hyperglycemia:
      • Diabetes mellitus: Type I diabetes mellitus is due to destruction of β cells in pancreatic islets (this is thought to be immune-mediated). Diabetes mellitus appears to be inherited in Keeshonds, with a possible familial form in Samoyeds. Diabetes mellitus has been associated with bovine virus diarrhea infection in cattle and paramyxovirus infection in llamas, through destruction of pancreatic islets. Type II diabetes mellitus is due to insulin resistance and animals do not necessarily need insulin to prevent hyperglycemia (called non-insulin dependent). Cats are prone to non-insulin dependent diabetes mellitus. This is thought to be associated with deposition of pancreatic amyloid (from amylin or amyloid polypeptide that is normally produced in β cells in the pancreas), which is related to pancreatic islet dysfunction. When islet destruction is widespread, cats do become insulin-dependent. 
      • Hyperadrenocorticism: In dogs with Cushing’s disease, hyperglycemia is due to insulin resistance from excess corticosteroids. Dogs with diabetes mellitus that become insulin resistant or require increasing doses of insulin, should be tested for underlying Cushing’s disease. Horses with pituitary par intermedia dysfunction (PPID) usually have pituitary adenomas that results in secretion of ACTH and other hormones that increase cortisol (see below). 
      • Acromegaly: Hyperglycemia is due to insulin resistance from high concentrations of growth hormone.
      • Hyperglucagonemia: Hyperglycemia is due to insulin resistance from high concentrations of glucagon, e.g. glucagon-secreting tumors (glucagonoma).
      • Hyperpituitarism/pituitary par intermedia dysfunction (PPID) in horses: Tumors in the pituitary gland can cause hyperglycemia through excess production of growth hormone or ACTH. Adenoma of the pituitary gland are the most common cause of PPID in horses. Horses with PPID are often hyperglycemia, which is usually mild (56% of horses in one study by Rohrbach et al., 2013 were hyperglycemic). The hyperglycemia is attributed to insulin resistance from hypercortisolemia caused by increased secretion of propriomelanocortin (POMC) peptides from the pituitary. In contrast, horses with metabolic syndrome are typically not hyperglycemic unless there are complicating diseases, causing a concurrent stress hyperglycemia. Horses with PPID tend to be older (>15 years, more than 80% in the study by Rohrbach), but often present with hirsutism and hyperhydrosis. Laminitis can be seen and horses tend to be underweight with abnormally distributed fat in the abdominal region (in contrast to horses with equine metabolic syndrome, which are usually obese). Due to the hypercortisolemia, they are predisposed to recurrent infections. A low-dose dexamethasone suppression test, ACTH stimulation (however, several studies have found that horses with PPID do not respond in an exaggerated manner), or a TRH stimulation test (the hyperplastic cells in the pituitary respond to TRH) can be helpful in diagnosing PPID in horses.
      • Pheochromocytomas: Some animals with these catecholamine-producing tumors are hyperglycemic.
    • Transient hyperglycemia
      • Hyperthyroidism in cats: Increases in glucose are often transient. The exact mechanism is unknown (? defective insulin secretion, ? enhanced sensitivity to catecholamines).
      • Acute pancreatitis: Hyperglycemia, which is usually transient, may occur due to stress, glucagon secretion and decreased insulin production.
      • Sepsis: Some animals with acute sepsis are hyperglycemic. This could be secondary to concurrent corticosteroid or catecholamine secretion (“stress”) or inflammatory cytokines, such as TNF-α (presumably due to insulin resistance [Hillebrand et all 2012]). The latter is thought to be the main mechanism by which sepsis, particularly acute, causes a hyperglycemia. However, with severe sepsis or sepsis of a longer duration, hypoglycemia may ensue.
      • Proximal duodenal obstruction: Adult cattle with proximal duodenal obstruction can have severe hyperglycemia (>500 mg/dL), accompanied by a severe hypochloremic metabolic alkalosis (sequestration of hydrochloric acid in the rumen) (Garry et al 1988). This is not specific for this syndrome (and is certainly not seen in all affected animals [Vogel et al 2012]) and can be seen in cattle that are stressed and have abomasal volvulus, right or left displaced abomasum or other causes of decreased abomasal outflow. In our experience, cattle with severe hyperglycemia have a poor prognosis (TJ Divers, personal communication).
      • Hypersomolar syndrome in crias: Crias can develop a syndrome of hyperosmolality that is characterized by marked hyperglycemia (usually > 300 mg/dL), hypernatremia, metabolic acidosis, and azotemia. This is thought to be due to a combination of stress and inadequate insulin response (camelids are relatively resistant to insulin,  do not release insulin as well as other species in response to hyperglycemia resulting in decreased glucose clearance from blood). Affected animals present with anorexia, lethargy, dehydration, fever and neurologic deficits (attributed to hyperosmolality) including wide-based stance, ataxia, seizures, and obtundation. Affected animals can have concurrent hypertriglyceridemia and high NEFAs (likely stress-induced lipolysis with insulin resistance, leading to increased VLDLs) (Cebra 2000, Buchheit et al 2010). Affected crias can respond to insulin infusion (Buchheit et al 2010).


Lack of glucose produces seizures as the brain relies entirely on glucose for its energy source. Neonatal animals are predisposed to hypoglycemia due to immature hepatic gluconeogenic pathways, low fat stores and muscle mass and rapid glycogen depletion. Hypoglycemia can be due to decreased production (e.g. inherited disorders, liver disease) or increased use (e.g. insulinomas, sepsis). The most common cause of hypoglycemia is an artifact. Common pathophysiologic causes are sepsis and, in dogs, hypoadrenocorticism and insulinoma. In cattle, lactation used to be a common cause of hypoglycemia (leading to type I bovine ketosis or clinical ketosis) but this is uncommon on modern dairy farms (rather type II or subclinical ketosis from excessive negative energy balance is now more common).

  • Artifact: False decreases in glucose concentration should be suspected in samples in which the glucose concentration is very low but the patient is not demonstrating clinical signs of hypoglycemia (e.g. seizures, disorientation).
    • Improper sample handling: Not separating serum from cells, e.g., mailing serum on clot. This artifact still occurs, albeit to a lesser extent, with corvac tubes, in which the serum is not removed from the tube. The silicon barrier does not always prevent glucose consumption, particularly when it is thin or breached (old tubes). Along with high potassium (in horses, some breeds of cattle, pigs, camelids, and some Asian breeds of dogs), a false decrease in glucose from improper sample handling is the most common artifact we see in mailed in samples (when there is a delay between separation of serum or plasma from cells or the serum or plasma is not separated from cells).
    • Bacteria in the sample: This can occur with bacterial contamination of the sample or infection. False decreases in glucose concentration have been reported in animals with high blood burdens of the hemotrophic bacteria, Mycoplasma (Burkhard and Garry, 2004).
  • Iatrogenic: Insulin administration.
  • Pathophysiologic: Pathophysiologic hypoglycemia results from decreased production of glucose, increased utilization of glucose by tissues, or in some cases both. Clinical signs associated with hypoglycemia may include lethargy, lack of coordination, exercise intolerance, polydipsia, blindness, convulsions, seizures, and coma (if glucose levels are severely low). 
    • Decreased production: Decreased production of glucose by the liver can occur secondary to inherited defects in gluconeogenic or glycogenolytic enzyme pathways, decreased stores of glycogen, decreased intake of glucose or liver disease (since this is the main site of glucose production).
      • Inherited defects: Hypoglycemia is a feature of certain glycogen storage diseases, namely deficiencies of α 1-4 glucosidase (Pompe disease) and glucose-6-phosphatase (von Gierke’s disease). This results in hypoglycemia from defective glycogenolysis.
      • Idiopathic: Juvenile hypoglycemia (usually affects toy and small breed dogs). This is thought to be due to hepatic immaturity, low liver stores of glycogen and insufficient gluconeogenesis to meet demands.
      • Liver disease: Severe liver disease and portosystemic shunts can produce hypoglycemia, however this is uncommon, particularly with shunts. More than 70% of the functional liver mass must be lost before hypoglycemia ensues.
    • Decreased intake: Starvation, malabsorption. In horses, glucose decreases if they are fed a high grain diet, with little roughage. Neonates are more susceptible to starvation-associated hypoglycemia, particularly dogs, because of low glycogen stores in the liver, little fat stores, and inefficient gluconeogenesis. Apparently, calves, foals and lambs are less susceptible to this condition (stated in Kaneko).
    • Increased use: This can be mediated by insulin release, decreased insulin antagonism (decrease of counter-regulatory hormones) or increased glucose use by tissues. Disorders that cause increased use can also decrease production (downregulation of gluconeogenesis, e.g. hypoadrenocorticism).
      • Idiopathic: Hypoglycemia of hunting dogs and endurance horses. A hypoglycemic state is reached from an imbalance in which glucose consumption (glycolysis) occurs at a much faster rate than glucose replenishment (gluconeogenesis and glycogenolysis).
      • Increased insulin secretion: Insulin-secreting tumors (insulinoma) or tumors secreting insulin-like growth factors (mesenchymal tumors such as leiomyoma, leiomyosarcoma, hepatic and renal tumors). Xylitol, an artificial sweetener, can cause hypoglycemia, through stimulating insulin release. This occurs mostly in the dog and can occur quite rapidly (within 30 minutes) of ingestion  (Zia et al., 2009). Xylitol can also cause hepatic failure in dogs and was associated with increased liver enzymes, hyperbilirubinemia, coagulation defects and hypoglycemia in one report (Dunayer and Gwaltney-Brant, 2006).
      • Decreased counter-regulatory hormones: This can occur with hypopituitarism (growth hormone or ACTH deficiency) or Addison’s disease (hypoadrenocorticism).
        • Hypoadrenocorticism: Hypoglycemia occurs due to decreased gluconeogenesis and increased insulin-mediated glucose uptake by skeletal muscle.
      • Sepsis: Hypoglycemia occurs due to liver dysfunction (defective gluconeogenesis and glycogenolysis), impairment of insulin degradation and enhanced glucose use secondary to endotoxemia. A true in vivo hypoglycemia secondary to consumption of glucose by the actual organism has been identified rarely in some animals with severe hemotropic Mycoplasma infections (e.g. Mycoplasma haemolamae; although artifactual hypoglycemia is more common). Studies in sick calves (<1 month of age) and foals (<1 week of age) show that hypoglycemia is a negative prognostic indicator, likely due to its association with sepsis (Hollis et al 2008, Treftz et al 2016, Treftz et al 2017
      • Bovine ketosis (type I) and ovine pregnancy toxemia (lactational hypoglycemia): During pregnancy, there are increased glucose demands from the fetus and the mammary glands (plasma glucose is the source of lactose). Ruminants are predisposed to hypoglycemia in late pregnancy or early lactation as they rely on gluconeogenesis for glucose production. Bovine type I ketosis results in anorexia, depression, decreased mild production, ketonemia, ketolactia, ketonuria and hypoglycemia. It usually occurs in dairy cows in the first 1-2 months of lactation due to increased demands for glucose by the mammary gland. It is initiated by poor diets or inappetance from other diseases. Note, that type I bovine ketosis is uncommon in current intensive dairy production, where high producing dairy cows are fed total mixed rations. Type II bovine ketosis (subclinical ketosis) which is not associated with hypoglycemia is far more common than type I ketosis (although the latter can still occur). Beef cows can also develop ketosis, especially in the last 2 months of pregnancy, when carrying twins. Ovine pregnancy toxemia occurs in sheep, carrying more than 1 fetus, that are calorically deprived or stressed. They, like beef cows, develop fatty liver and ketoacidosis and may die of liver dysfunction. Ketosis has also been reported rarely in lactating dairy goats and dogs.
      • Exertional hypoglycemia: This has been identified in hunting dogs and endurance horses, where demand exceeds supply.


  • Kaneko JJ. Carbohydrate metabolism and its diseases. In: Kaneko JJ, Harvey JW, Bruss ML, eds. Clinical Biochemistry of Domestic Animals. 5th ed. Toronto: Academic Press, 1997, pp 45–81.
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