In the 1950’s, a group of entomologists at the University of Illinois (Carter et al., 1952) found that the successful growth of the yellow mealworm (Tenebrio molitor) in culture required the feeding of a natural substance that was found to be present in milk, yeast and many animal tissues. The substance was purified, named “vitamin Bt” and later identified as carnitine.
Carnitine is a vitamin-like substance that is a quaternary amine, beta-hydroxy-gamma-trimethylaminobutyrate. Carnitine can be synthesized in the body of animals through a metabolic process involving several cofactors (lysine and methionine), four vitamins (vitamin C, niacin, vitamin B12 and choline) and reduced iron. Carnitine is synthesized endogenously from lysine (carbon backbone) and methionine (methyl group donor). However, these amino acids are not directly converted to carnitine but are required for synthesis of the endogenous carnitine precursor trimethyllysine. The rate of protein turnover and the trimethyllysine content in proteins are the main factors determining the rate of endogenous carnitine synthesis (Vaz and Wanders, 2002). Despite increased hepatic carnitine concentration around calving (Grum et al., 1996), insufficient endogenous carnitine synthesis might contribute to fatty liver-development in periparturient dairy cows (Carlson et al., 2007b).
Deficiency in any cofactor will cause L-carnitine deficiency. In rats, total acid-soluble carnitine and free carnitine in plasma and tissues were reduced in a vitamin B6 deficiency but increased when vitamin B6 was provided in a repletion diet (Cho and LeKlem, 1990; Ha et al., 1994). It has been suggested that early features of scurvy (fatigue and weakness) may be attributed to carnitine deficiency. Vitamin C is a cofactor for two alpha-ketoglutarate-requiring dioxygenase reactions (epsilon-N-trimethyllysine hydroxylase and gamma-butyrobetaine hydroxylase) in the pathway of carnitine biosynthesis. Carnitine concentrations are variably low in some tissues of vitamin C-deficient guinea pigs (Rebouche, 1991).
Choline has also been shown to affect carnitine homeostasis in humans and guinea pigs (Daily and Sachan, 1995). Choline supplementation resulted in decreased urinary excretion of carnitine in young adult women. Choline resulted in the conservation of carnitine in guinea pigs. In adult women, Hongu and Sachan (2003) concluded that the choline-induced decrease in serum and urinary carnitine is buffered by carnitine preloading, and these supplements shift tissue partitioning of carnitine that favors fat mobilization, incomplete oxidation of fatty acids and disposal of their carbons in urine.
Carnitine is synthesized in part from methyl groups derived from choline and other donors via S-adenosylmethionine (Lehninger, 1975). Choline may affect lipid metabolism indirectly by influencing synthesis of carnitine. A series of studies with lactating dairy cows found that plasma and liver carnitine could be increased either by feeding or abomasal infusion of carnitine, but that milk yield and dry matter intake were not affected by carnitine supplementation (LaCount et al., 1995; 1996a, b). However, total tract digestibility of lipids and energy increased in response to carnitine supplementation by either route (LaCount et al., 1995).
Carnitine appears to be absorbed across the gut by an active process dependent on Na+ as well as by a passive diffusion that may be important for the absorption of large doses of the factor. The uptake of carnitine from the intestinal lumen into the mucosa is rapid, and about one-half of the carnitine taken up is acetylated in that tissue. Absorption of carnitine in dietary supplements (0.5 to 4 g per day) is 15% to 25% (Rebouche, 2006).
Tissues such as cardiac muscle and skeletal muscle require carnitine for normal fuel metabolism but cannot synthesize carnitine and are totally dependent on the transport of carnitine from other tissues. Free carnitine is excreted in urine, with the principal excretory product being trimethylamine oxide (Mitchell, 1978). Carnitine is highly conserved by the human kidney, which reabsorbs more than 90% of filtered carnitine, thus, playing an important role in the regulation of carnitine concentration in blood. For the dog, 95% to 98% of the carnitine body pool is in skeletal muscle and the heart (Rebouche and Ergel, 1983). With rats, Flores et al. (1996) reported that the small intestine is a considerable and previously unrecognized proportion of the carnitine pool of suckling animals.
Carnitine is required for transport of long-chain fatty acids into the matrix compartment of mitochondria from cytoplasm for subsequent oxidation by the fatty acid oxidase complex for energy production. The oxidation of long-chain fatty acids in animal tissues is dependent on carnitine because it allows long-chain acyl-CoA esters to cross the mitochondrial membrane, which is otherwise impermeable to CoA compounds. Carnitine facilitates the beta-oxidation of long-chain fatty acids in the mitochondria by transporting the substrate into the mitochondria. Carnitine acyltransferase is the enzyme responsible for this shuttle mechanism. It exists in two forms, carnitine acyltransferase I and carnitine acyltransferase II. After the long-chain fatty acid is activated to acyl-CoA, it is converted to acylcarnitine by the enzyme carnitine acyltransferase I and crosses to the matrix side of the inner mitochondrial membrane. Carnitine acyltransferase II then releases carnitine and the acyl-CoA into the mitochondrial matrix. Acyl-CoA is then catabolized via beta-oxidation (Borum, 1991). Thus, utilization of long-chain fatty acids as a fuel source depends on adequate concentrations of carnitine.
L-carnitine has several other functions such as altering the acetyl-CoA:CoA ratio, transporting medium-and short-chain fatty acids from peroxisomes to mitochondria, and modulating flux of intermediates through pathways associated with fatty acid, glucose, and nitrogen metabolism (Ji et al., 1996; Owen et al., 2001). The role of carnitine in hepatic fatty acid oxidation suggests that carnitine status might influence the degree of liver lipid accumulation in periparturient dairy cows. In dairy cows, carnitine stimulates hepatic fatty acid oxidation in vitro (Jesse et al., 1986; Drackley et al., 1991b). Prepartum increases in liver carnitine concentration were associated with decreased liver triglyceride accumulation during the transition period (Grum et al., 1996). It was shown previously that abomasal infusion of L-carnitine increased in vitro hepatic fatty acid oxidation regardless of amount of feed intake, and decreased liver lipid accumulation in feed-restricted cows (Carlson et al., 2006).
Another role of carnitine may be to protect cells against toxic accumulation of acyl-CoA compounds of either endogenous or exogenous origin by trapping such acyl groups as carnitine esters, which may then be transported to the liver for catabolism or to the kidney for excretion in the urine. Carnitine also has functions in other physiological processes critical to survival, such as lipolysis, thermogenesis, ketogenesis and possibly regulation of certain aspects of nitrogen metabolism (Borum, 1985).
Carnitine was found to be beneficial in detoxification of aflatoxins (Yatim and Sachan, 2001). For rats, there was a modification of aflatoxin binding to proteins and DNA by carnitine which reduced the carcinogenic and hepatotoxic potential of aflatoxins.
Higher animals, including mammals, can synthesize carnitine. However, recent studies have indicated that the biosynthesis of carnitine may be limited or inadequate in certain animals. The liver and kidney of mammals are able to synthesize carnitine from lysine and methionine; however, endogenous biosynthesis alone is not sufficient to keep carnitine concentrations at adequate levels (Duran et al., 1990).
Carnitine status in mammals is influenced by dietary intake, endogenous synthesis, and reabsorption of carnitine and carnitine precursors by the kidney (Rebouche and Seim, 1998). The rate of carnitine biosynthesis is largely dependent upon the supply of trimethyllysine from turnover of proteins containing this amino acid (Vaz and Wanders, 2002). Lysine (carbon backbone) and methionine (methyl group donor) are required for trimethyllysine synthesis. In the rat, several tissues are capable of converting trimethyllysine to the immediate carnitine precursor γ-butyrobetaine (Vaz and Wander, 2002). In most species only the liver possesses the enzyme necessary for converting γ-butyrobetaine to carnitine, although in some species the kidney, brain, and testis are capable of synthesizing carnitine from γ-butyrobetaine (Vaz et al., 1998; Galland et al., 1999).
For common livestock species, there are no established nutritional requirements for carnitine. Ruminant studies showed positive responses for supplemental carnitine; 1.5 g/day in beef steers (Greenwood et al., 2001) and 6 to 100 g/day for dairy cattle (Carlson et al., 2006; 2007a, b).
Carnitine has both D- and L-forms, but only the L-form is biologically active and occurs in nature. In general, foods of plant origin are low in carnitine, whereas animal-derived foods are rich in carnitine (Mitchell, 1978; Rebouche, 2006). Red meats and dairy products are particularly rich sources. The carnitine concentration increases in the order of fish, poultry, pork, beef and lamb. In general, the redder the meat, the higher the concentration of carnitine. Typical concentrations of carnitine could be 600 µg per kg (272.7 µg per lb) for beef, 45 to 90 µg per kg (20.5 to 40.9 µg per lb) for chicken and 75 µg per kg (34.1 µg per lb) for lamb (Mitchell, 1978). Carnitine is located principally in skeletal muscle, which has about 40 times the concentration of carnitine found in blood. On the contrary, grains such as barley, corn and wheat have undetectable or negligible concentrations. Most plant foods that are low in carnitine are also likely to be low in lysine and methionine, the precursors of carnitine. Milk is essential for the nursing mammal’s carnitine supply. Although carnitine is synthesized in growing young and adult animals, previous studies in humans provide evidence that exogenous carnitine is necessary to maintain normal fat metabolism during infancy. Studies by Davis (1989) indicated that up to 50% of tissue carnitine in suckling rats is derived from the mother’s milk. Studies with neonatal rabbits (Penn and Schmidt-Sommerfeld, 1988) and rats (Flores et al., 1996) demonstrated that body tissue concentrations of carnitine are greatly diminished in newborns deprived of milk during early life. Coffey et al. (1991) showed that diminished dietary intake is associated with decreased levels of carnitine in liver, but not heart or muscle, in neonatal piglets receiving low levels of dietary carnitine compared with piglets receiving carnitine supplementation. These observations indicate the importance of milk. The demand for carnitine during the suckling period may exceed the capacity for its synthesis.
Carnitine is commercially available to the feed industry as a 50% L-carnitine product. Carnitine is very hygroscopic and easily soluble in water and has a molecular weight of 161.2.
In carnitine deficiency, fatty acid oxidation is reduced, and fatty acids are diverted into triglyceride synthesis, particularly in the liver. Mitochondrial failure develops in carnitine deficiency when there is insufficient tissue carnitine available to buffer toxic acyl-coenzyme (CoA) metabolites. Toxic amounts of acyl-CoA impair the citrate cycle, gluconeogenesis, the urea cycle and fatty acid oxidation. Carnitine replacement induces excretion of toxic acyl groups in the urine (Stumpf et al., 1985). Skeletal muscles are generally involved, with weakness, lipid myopathy and myoglobinuria often aggravated or precipitated by fasting or exercise. For exercising pigeons, L-carnitine supplementation improved fatty acid oxidation efficiency during heavy exercise (Janssens et al., 1998).
If carnitine deficiency involves the liver, the supply of ketones and the utilization of long-chain fatty acids during starvation are cut off; all tissues become glucose dependent. When liver carnitine is depleted, starvation tends to cause nonketotic, insulinopenic hypoglycemia. Because liver hepatocytes depend on fatty acids for their energy requirements during fasting, carnitine depletion may also cause clinical liver dysfunction, shown by hyperammonemia, encephalopathy and hyperbilirubinemia (Feller and Rudman, 1988). Skeletal muscles are generally involved; for monogastric species clinical signs include weakness, lipid myopathy and myoglobinuria often aggravated or precipitated by fasting or exercise. The heart, like skeletal muscle, is dependent on fatty acids for energy during fasting, and heart failure and arrhythmias are frequent manifestations of systemic carnitine deficiency. The heart derives approximately 60% of its ATP supply from beta-oxidation of fatty acids. Carnitine concentrations in the heart are normally very high in many species (Rebouche and Paulson, 1986).
Supplemental carnitine has been administered to prepartum and lactating dairy cows (Carlson et al., 2006; 2007a, b) and to growing steers (Greenwood et al., 2001). For dairy cows, the objectives were to determine the effects of dietary carnitine on liver fat accumulation, hepatic lipid and carbohydrate metabolism, tissue carnitine concentrations, and lactation performance of Holstein cows during the periparturient period. Milk carnitine concentrations were elevated by all amounts of carnitine supplementation. The 50 or 100 g per day carnitine treatments increased milk fat concentration, although milk fat yield was unaffected. Liver and muscle carnitine concentrations were markedly increased by supplemental carnitine.
All carnitine treatments decreased liver total lipid and triacylglycerol accumulation on day 10 after calving. In addition, carnitine-supplemented cows had higher liver glycogen during early lactation. In general, carnitine supplementation increased in vitro palmitate β-oxidation by liver slices. Carnitine effectively decreased liver lipid accumulation as a result of greater capacity for hepatic fatty acid oxidation. By decreasing liver lipid accumulation and stimulating hepatic glucose output, carnitine supplementation would improve glucose status and diminish the risk of developing metabolic disorders during early lactation.
Greenwood et al. (2001) evaluated the effects of supplemental carnitine on protein deposition and metabolism of growing cattle and the performance of finishing cattle. Steers receiving carnitine tended to have fatter carcasses, as indicated by tendencies for thicker backfat, higher marbling scores, and higher yield grades. Although carnitine supplementation did not alter lean deposition in growing steers, it did alter plasma non-esterfied fatty acids (NEFA) concentrations of growing steers fed a corn-based diet and also seemed to increase fat deposition in finishing cattle.
Only the physiological L-carnitine should be used for fortification of diets. Supplementation of 50 to 500 mg per kg (22.7 to 227 mg per lb) of diet have been used as supplementation doses for various livestock species. Although carnitine has been studied in humans and under laboratory conditions for many years, its effectiveness in promoting the performance and well-being of domestic animals has only recently received attention. A role for carnitine in swine and fish diets and tentatively for poultry has been established and continued research may find that it has a place in the production of ruminant livestock species.
The periparturient period is a critical time in the lactation cycle. Dairy cows are susceptible to a number of metabolic disorders and infectious diseases during this period (Goff, 2000). An increased understanding of lipid metabolism, in particular fatty acid oxidation, may allow the development of nutritional and management approaches to prevent the development of metabolic disorders, such as hepatic lipidosis and ketosis, in dairy cows (Dam and Drackley, 2005).
The transition from gestation to lactation generally is accompanied by decreased dry matter-intake, negative energy balance, and extensive mobilization of body tissue stores of nutrients (Drackley, 1999). Fatty liver occurs when excessive amounts of NEFA are esterified to triacylglycerol (TG) as a result of suboptimal hepatic fatty acid oxidation and limited TG export. Fatty liver is associated with an increased risk of ketosis, displaced abomasum, metritis, immune suppression, and decreased reproductive performance (Bobe et al., 2004). Researchers have attempted to alter hepatic metabolism and prevent fatty liver by reducing NEFA mobilization from adipose tissue, by enhancing hepatic fatty acid oxidation, or by increasing TG export from the liver through manipulation of pre or postpartum nutrition (Bobe et al., 2004).
Carnitine is a required cofactor for carnitine palmitoyltransferase-I (CPT-I), which condenses carnitine with activated long-chain fatty acids (LCFA) for transport from cytosol into mitochondria, therefore, carnitine is essential for mitochondrial oxidation of LCFA. Hepatic β-oxidation of LCFA is stimulated by exogenous carnitine in several species (Drackley et al., 1991a, b; Owen et al., 2001; Spanio et al., 2003). Additionally, carnitine supports β-oxidation by transporting short-and medium-chain fatty acids from peroxiomes to mitochondria and by altering the intramitochondrial ratio of acetyl-coenzyme A (Rebouche and Seim, 1998). Further, as a result of enhanced LCFA β-oxidation, carnitine supplementation has been shown to increase hepatic glucose production by stimulating the flux of metabolites through pyruvate carboxylase (Owen et al., 2001). Adaptations in hepatic lipid and carbohydrate metabolism are required for optimal health and productivity of dairy cows. The rate and extent of adipose tissue lipolysis and the capacity for hepatic LCFA β-oxidation might be the most important factors regulating the degree of liver TG accumulation (Drackley and Andersen, 2006). The metabolic functions of carnitine suggest that carnitine supply would be important during the periparturient period. Recent research (Carlson et al., 2006; 2007a, b) with transitional dairy cows has shown that carnitine administration decreased liver fat as a result of greater liver fatty acid oxidation, improved glucose status and reduced the risk of metabolic disorders during early lactation.
Little research has been conducted with carnitine and growing beef cattle, and results for beef cattle fed high-grain diets have been less than conclusive. Heifers fed a high-grain diet had lower quality grades when they received 1 g/d carnitine (Hill et al., 1995b), whereas steers and heifers receiving 100 ppm carnitine in a high-grain diet had higher quality meat grades (Hill et al., 1995a).
Livestock and poultry studies to determine maximum tolerance for carnitine are lacking. For human patients, carnitine supplementation at dosages that far exceed the usual dietary intake of carnitine have been administered (Goa and Brogden, 1987). Oral dosages of 100 mg per kg (45.5 mg per lb) have been given to infants and 1 to 3 g daily to adult humans with little problem. Some patients experienced diarrhea, but not if they started with smaller dosages and then increased gradually (Borum, 1991).
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