This paper discusses animal testing to diagnose mineral deficiencies and provides a summarization of the most commonly utilized tissues and fluids used for diagnosing mineral deficiencies.
Many minerals are essential for optimal growth, biological function, and productivity in ruminants. Historically, mineral testing has been performed on diets and/or feedstuffs to ensure adequate dietary concentrations. However, general mineral analysis does not identify chemical form of the minerals, which can alter bioavailability and utilization.
Mineral deficiencies are best diagnosed by testing for biological compounds or enzymes dependent upon the particular mineral. This type of testing is often impractical on the farm, due to costs or rigorous sample handling requirements. But, when possible, it will eliminate the need to know dietary mineral bioavailability and/or whether potentially antagonistic minerals are present. For minerals that do not have identified physiologic indices, direct quantification from tissues or serum may provide a reliable indication of adequacy or deficiency.
Mineral deficiencies may be suggested by clinical disease or tissue lesions. However, proof often requires analytical verification since most mineral deficiencies do not have very unique clinical signs or lesions. In some instances, circumstantial proof of a deficiency can be provided by positive response to supplementation of the suspect mineral. But, the positive response may have nothing to do with the supplementation; it may only be a response to correction of some other clinical condition.
An individual mineral may have multiple means of measurement for identification of deficiencies, but most have one that is more specific than the others. For example, dietary concentrations may or may not be reflective of the amount of bioavailable minerals. Or, an individual tissue concentration may or may not reflect functionally available mineral concentrations at the target or functional site.
The age of the animal being tested also is important for proper interpretation of mineral status. For example, feti accumulate some minerals at different rates during gestation, necessitating adequate aging of the fetus for interpretation. In addition, some minerals, for which little is provided in milk, accumulate at higher concentrations during gestation in order to provide the neonate with adequate body reserves for survival until it begins grazing. This is especially prevalent with copper, iron, selenium, and zinc. Thus, the normal range for these minerals would be higher in early neonates than in an adult animal.
When individual animals are tested, the prior health status must be considered in interpreting mineral concentration of tissues. Disease states can shift mineral from tissues to serum or serum to tissues. For example, diarrhea can result in significant loss of sodium, potassium, and calcium from the body. Acidosis will cause electrolyte shifts between tissues and circulating blood. It is known that infectious disease, stress, fever, endocrine dysfunction, and trauma can alter both tissue and circulating serum/blood concentrations of certain minerals and electrolytes. Thus, multiple animal evaluations are much more reflective of mineral status within a group than testing individual animals that are ill or have died from other disease conditions.
This paper discusses animal testing to diagnose mineral deficiencies and provides a summarization of the most commonly utilized tissues and fluids used for diagnosing mineral deficiencies.
Live Animal Sampling
The most common samples from live animals are serum and whole blood. These samples are adequate for several minerals; but it must be recognized that some disease states, as well as feeding times, can result in serum fluctuations. Other common samples from live animals include liver biopsies, urine, and milk. Since milk mineral content can vary through lactation, vary across lactations, and be affected by disease; it is not typically used to identify deficiencies. Serum should be separated from the red/white blood cell clot within 1 to 2 hours of collection. If the serum sets on the clot for long periods of time, minerals that have higher intracellular content than serum can leach into the serum and falsely increase the serum content. Minerals for which this commonly occurs include potassium and zinc. In addition, hemolysis from both natural disease and poor collection technique can result in increased serum concentrations of iron, manganese, potassium, selenium, and zinc.
The best type of tube for collecting serum or whole blood is royal blue-top vaccutainer tubes, as they are trace-metal free. Typical red-top clot tubes will give abnormal results for zinc content as a zinc containing lubricant is used on the rubber stoppers. For minerals other than zinc, serum samples from the typical red-top clot tubes are adequate. Samples should be appropriately stored for preservation. Liver biopsies, urine, and serum can be stored frozen long term or refrigerated if analysis is to be completed within 1-2 weeks. Whole blood and milk should be refrigerated, but not frozen, as cell lysis or coagulation of solids will result.
Post-Mortem Animal Sampling
A variety of post-mortem animal samples can be analyzed for mineral content. The most common tissue analyzed for mineral content is liver, as it is the primary storage organ for many essential minerals. In addition, bone is used as the primary storage organ for calcium, phosphorous, and magnesium. Other post-mortem samples that can be used for diagnosing mineral deficiencies include urine and ocular fluid. Post mortem samples should be stored frozen until analyzed to prevent tissue degradation. If samples are to be analyzed within 1-2 days, they can be stored under refrigerated conditions.
Analysis for calcium deficiency falls into two distinct classes. The first is metabolic calcium deficiency, or milk fever. The second is a true nutritional deficiency associated with long term dietary calcium deficits.
Metabolic calcium deficiency is diagnosed by a low systemic or circulating calcium level. In live animals, serum is used to determine circulating calcium content. However, testing is more difficult in dead animals; as serum collected postmortem will not accurately reflect serum calcium levels prior to death. However, circulating serum calcium content can be approximated from analysis of ocular fluid, with a vitreous to serum ratio of approximately 0.54 (McLaughlin and McLaughlin, 1987).
True nutritional calcium deficiency is associated with weak, unthrifty animals that have swollen joints, lameness, weak bones, and a propensity for broken bones (Puls, 1994). Analytical verification of calcium deficiency requires bone analysis, since approximately 98-99% of the body calcium content is in bone and serum concentrations are maintained by both diet and turnover of bone matrix. The bone analysis should be reported on a fat-free, dry weight basis to remove the age variability of moisture and fat content.
Cobalt deficiency is associated with deficiency of Vitamin B12 (cobalamin) in ruminants. Deficiency is associated with decreased feed intake, poor feed conversion, reduced growth, weight loss, hepatic lipidosis, anemia, immunosuppression, and impaired reproductive function (Graham, 1991; Puls, 1994). Cobalt deficiency can also lead to subnormal copper retention in the liver.
Tissue and serum concentrations of cobalt are generally quite low, as B12 is produced in the rumen by the microflora. Since cobalt concentrations may not truly reflect the B12 concentrations, cobalt deficiency is best determined by serum or liver Vitamin B12 levels. However, the numerous forms of cobalamins produced by ruminants differ in bioactivity which makes interpretation of analytical results difficult (Mills, 1987). Cobalamin is absorbed into circulation and small amounts are stored in the liver. Liver cobalt concentration best reflects the animal’s overall status, but it may not be truly reflective of vitamin B12 status.
Copper deficiency is a common nutritional problem in ruminants, but copper excess is also commonly encountered, especially in sheep. Clinical signs of deficiency cover a large array of adverse effects (Graham, 1991; Puls, 1994). Reduced growth rates, decreased feed conversion, abomasal ulcers, lameness, poor immune function, sudden death, achromotrichia, and impaired reproductive function are commonly encountered with copper deficiency.
Analyzing liver tissue is good for diagnosing copper deficiency, although much testing is performed on serum. Deficiency within a herd will result in some animals that have low serum copper concentrations, but serum content does not decrease until liver copper stores are fairly depleted. In herds that have tested liver and found a high incidence of deficiency, it was not uncommon for a high percentage of the animals to have normal serum concentrations. The Utah Veterinary Diagnostic Laboratory commonly recommends that 10% of a herd or a minimum of 10-15 animals be tested to have a reasonably high probability of diagnosing copper deficiency with serum analysis. Even with a severely deficient herd, low serum copper concentrations may be seen in as few as 20% of the individuals tested. Herds classified as marginally deficient based on liver testing may have predominantly normal serum copper concentrations. Thus, serum copper analysis should be viewed as a screening method only. Another factor that can influence diagnosis of copper deficiency in serum is the presence of high serum molybdenum. As the copper-sulfur-molybdenum complex that forms is not physiologically available for tissue use, normal serum copper content in the presence of high serum molybdenum should always be considered suspect.
Copper deficiency can also be diagnosed via analysis of copper containing enzymes. The two most common enzymes used are ceruloplasmin and superoxide dismutase (Mills, 1987; Suttle, 1986). Low concentrations of these enzymes in serum and whole blood, respectively, are diagnostic for copper deficiency. But, ceruloplasmin concentrations can increase with inflammatory disease. The higher cost of enzyme analyses compared to liver copper analysis, often limits their utilization.
Iron is essential for the normal cellular function of all cell types. Iron deficiency is associated with reduced growth, poor immune function, weakness, and anemia (Graham, 1991; Puls, 1994). Although offspring are typically born with liver reserves of iron (provided the mother had adequate iron reserves), milk has a low iron content. This leads to iron deficiency over time in animals fed a milk-only diet. Both liver and serum concentrations are commonly utilized to diagnose iron deficiency.
When using serum to measure iron content, samples that have evidence of hemolysis should not be used, since the iron content will by increased by the ruptured red blood cells. In addition, disease states can alter serum and liver iron concentrations as the body both tries to limit availability of iron to growing organisms and increases the availability of iron to the body’s immune cells. Thus, interpretation of iron status should be made with consideration of the overall health of the animal. Other factors that can be used to assist with diagnosis of iron status include: serum iron binding capacity, serum iron binding saturation, red blood cell count, packed cell volume, serum hemoglobin concentration, and ferritin concentration (Smith, 1989). But a variety of clinical conditions can cause these values to vary, including bacterial infections, viral infections, other types of inflammation, hemorrhage, bleeding disorders, and immune mediated disorders.
Similar to calcium, analysis for magnesium deficiency falls into two distinct classes; metabolic magnesium deficiency (grass tetany) and true nutritional deficiency due to long term dietary magnesium deficits.
Analysis for metabolic magnesium deficiency depends on detection of low systemic or circulating magnesium levels. In live animals, serum is tested to determine circulating magnesium content. But, ruminants with recumbency or tetany may have normal serum magnesium, as tissue damage that occurs releases magnesium into the serum from the soft tissues. In dead animals testing is more difficult, as serum collected post-mortem will not accurately reflect true serum magnesium content prior to death. Circulating serum magnesium content can be approximated from analysis of ocular fluid, with a vitreous to serum ratio of 1.05 (McLaughlin and McLaughlin, 1987). Urine is another post-mortem sample that can be analyzed, since the kidneys minimize magnesium loss in the urine when serum magnesium is low.
True nutritional magnesium deficiency is not recognized in ruminants, except under experimental conditions. This syndrome is associated with weak, unthrifty animals that have weak bones, low bone ash, and calcification of soft tissues. Analytical verification of true magnesium deficiency requires bone analysis, since approximately 70% of body magnesium is in bone. The bone analysis should be reported as fat-free, dry weight to remove the age variability of moisture and fat content.
Manganese deficiency in ruminants is associated with poor reproductive function, skeletal abnormalities in calves, and less than optimal productivity (Graham, 1991; Puls, 1994). Cystic ovaries, silent heat, reduced conception rates, and abortions are typical reproductive effects. Manganese deficient calves can be weak, small, and develop enlarged joints or limb deformities. Manganese deficiency, although not reported often, is identified routinely in dairy cattle when tested. Most beef cattle show normal manganese concentrations in liver, blood, and serum, but in these same matrices greater than 50%, 75%, and 95%, respectively, of dairy cattle tested are below recommended normal concentrations (unpublished data). This may, in part, be due to high calcium and phosphorous content of dairy rations, which can reduce bioavailability of manganese.
Of the samples available, liver is the most indicative of whole body status, followed by whole blood then serum. As red blood cells have higher manganese content than serum, hemolysis can result in increased serum content. Since the normal serum concentration of manganese is quite low, many laboratories do not offer this analysis because of inadequate sensitivity. Overall, response to supplementation has frequently been used as a means of verifying manganese deficiency, but it is critical that a bioavailable form be utilized.
Phosphorous deficiency is somewhat difficult to measure in animal tissues. Serum and urine phosphorous concentrations can aid in diagnosing deficiency, but mobilization of bone phosphorous acts to maintain serum phosphorus content. Therefore, significant drops in serum and urine phosphorus may take weeks to develop. Serum phosphorous should be measured as inorganic phosphorous to aid in interpretation. Longer term phosphorous deficiency can be diagnosed post-mortem by measuring bone or bone ash phosphorous content. Dietary phosphorous and/or response to supplementation are better indicators of deficiency than tissue concentrations unless severe long term deficiency has occurred. The predominant effects of low dietary phosphorous are associated with diminished appetite and its resultant effects. Depressed feed intake, poor growth, and weight loss are common with phosphorous deficient diets. Longer term phosphorous deficiency results in poor reproductive performance, impaired immune function, bone abnormalities, and pica.
Tissue concentrations of potassium poorly correlate with dietary deficiency. Of the animal samples available, serum potassium is the best indicator of deficiency; but disease states can cause electrolyte shifts that result in lowered serum potassium when dietary deficiency has not occurred. In addition, serum that is hemolyzed or left on the clot too long may have falsely increased potassium content due to loss from the red blood cells. Thus, dietary potassium concentrations are a better guide to potassium deficiency. Dietary potassium deficiency affects intake, productivity, heart function, and muscle function. Common clinical signs of severe potassium deficiency include: diminished feed intake, reduced water intake, pica, poor productivity, weakness, and recumbency.
Selenium is commonly identified as deficient in ruminants. Selenium deficiency in ruminants is associated with adverse effects on growth, reproduction, immune system function, offspring, and muscle tissues (Graham, 1991; Puls, 1994). White muscle disease, a necrosis and scaring of cardiac and/or skeletal muscle, is linked to severe selenium deficiency; although it can be caused by vitamin E deficiency as well. Poor growth rates, immune function, and reproductive performance can be observed with less severe selenium deficiency.
Diagnosis of a deficiency can be made by analysis of liver, whole blood, or serum for selenium content or by analysis of whole blood for glutathione peroxidase, a selenium dependent enzyme (Ullrey, 1987). The most specific analysis is that of whole blood glutathione peroxidase, as it verifies true functional selenium status. Liver is the optimal tissue to analyze for selenium content as it is a primary storage tissue. With serum and whole blood, the former better reflects recent intake, while the latter better reflects long term status. Since glutathione peroxidase is incorporated into the red blood cells when they are made and the cells have a long half-life, selenium content is a reflection of intake over the previous months. In order to adequately diagnose selenium deficiency, the dietary form of selenium in the diet is important. Natural selenium, predominantly in the form of selenomethionine, is metabolized and incorporated into selenium dependent proteins, but can also be incorporated into non-specific proteins in place of methionine. Inorganic selenium is metabolized and only incorporated into selenium dependent proteins. Thus, normal concentrations in serum and whole blood differ, depending on whether the dietary selenium is a natural organic form or an inorganic supplement.
Tissue concentrations of sodium poorly correlate with dietary deficiency. Serum and urine are the best for measuring sodium deficiency; but disease states can cause electrolyte shifts that result in lowered serum or urinary sodium even when dietary concentrations are adequate. Thus, dietary sodium concentrations are a better guide to diagnosing a deficiency. Dietary sodium deficiency affects feed intake and productivity. Common clinical signs of severe sodium deficiency include: diminished feed intake, decreased water intake, poor productivity, and pica.
Zinc is an essential mineral that is required by all animal cells. Zinc plays a role in numerous enzymatic reactions (Graham, 1991; Puls, 1994). Deficiencies of zinc are associated with poor growth, immune function, reproductive performance, and offspring viability; as well as skin lesions in severe cases. Tissue zinc concentrations do not reflect body status well (Mills, 1987). Liver and serum are the best indicators of zinc status. But, serum and liver zinc can be altered by age, infectious diseases, trauma, fever, and stress. It has been suggested that pancreatic zinc content is the best means of truly identifying zinc deficiency. Response to zinc supplementation has shown that some animals having low-end normal liver or serum zinc can still show improvement in some clinical conditions. Thus, liver and serum only verify deficiency when these samples have a very low zinc content.
A variety of samples can be tested for mineral content, but may not provide any indication of the overall mineral status of the animal. Appropriate diagnosis of mineral deficiencies involves good evaluation of groups of animals. This evaluation should include a thorough health history, feeding history, supplementation history, and analysis of several animals for their mineral status.
Dietary mineral evaluation should augment the mineral evaluation of animal groups. If minerals are deemed to be adequate in the diet, but the animals are found to be deficient, antagonistic interactive effects of other minerals need to be investigated. As an example, high sulfur or iron can cause deficiencies in copper and selenium even when there are adequate concentrations in the diet.
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