Introduction
The circulating concentrations of glucose are very high in both wild and domestic birds compared to mammals when animals of similar body mass are compared (
Braun and Sweazea, 2008) (Table 1 and Fig. 1). Allometric analysis of a large database (
n = 97 for wild birds and
n = 162 for mammals) shows parallel lines with identical slopes when plasma glucose concentrations are plotted against the log of body mass:
In birds, circulating concentration of glucose (mM) = 15.3 - 0.44 log body mass (kg).
In mammals, circulating concentration of glucose= 7.6 - 0.44 log body mass (kg).
The y intercept of the two linesis 15.3 mM (or 275 mg/dL) in wild birds and 7.6 mM (137 mg/dL) in mammals (
Braun and Sweazea, 2008); differing by a factor of two. Thus, non-domestic birds have plasma glucose concentrations twice as high as that of mammals. When individual speciesare compared, it is apparent that some birds have plasma glucose levels that are 6 fold higher that mammals of similar body mass. It has been suggested that as birds are less long-lived than mammals, the relatively high plasma glucose concentrations are of little consequence to the pathologies of these animals. A simple empirical observation of a 20 gm body mass mouse and a 20 gm parakeet belies this suggestion; the mouse is senescent at 18 - 24 months and the parakeet will live 20 to 20 five years.
Glucose excretion by birds
Under normal glucose homeostasis, little or no glucose appears in the uretheral urine of birds. Although it should be noted that when the excreta of birds are collected, small quantities of glucose will be present as result of bacterial fermentation in the lower gastrointestinal tract. It is of interest that no glucose escapes in the uretheral urine of birds considering the high plasma concentration and the glomerular filtration rates (GFRs) that, on an allometric basis, do not differ significantly from those of mammals (
Yokota et al., 1985). Plasma concentration of glucose multiplied by the GFR yields a parameter called the filtered load (FL) of glucose. This is a much higher value for birds given the high plasma glucose levels. Yet no glucose appears in the urine. It has been determined and quite well characterized for mammal kidneys that there are two glucose transport proteins (sodium/glucose co-transport proteins, SGLTs) present in the proximal renal tubule that, under normal glucose homeostasis, reabsorb all the glucose that is filtered at the glomerulus (
Turk and Wright, 1997). For birds, all that is known at this time is that no glucose appears in the urine. Therefore, some isoform of the SGLTs must be present within the avian kidney. SGLT1 appears in the chicken genome, but it is the protein that is present in the lower gastrointestinal tract. To date, little or no work has been conducted on the glucose transport capacity of the avian kidney. This is an area that warrants attention.
Control of carbohydrate metabolism in birds
There is a marked tendency for researchers to consider birds either as wild wilds or one of the domesticated species or poultry. There might to be changes in the physiologic control mechanisms due to domestication and/or to intensive selective breeding either for laying or for rapid growth. Chickens were developed from the ancestral Red Jungle Fowl (
West and Zhou, 1988). Comparative genomics of chickens with the ancestral Red Jungle Fowl has identified deletions (
Rubin et al., 2010). The available evidence (discussed below) supports close consistency in the control of metabolism in wild and domesticated birds.
Pancreatic hormones and metabolism
Glucagon
Glucagon is generally considered as the major hormone controlling metabolism in birds. There have been extensive studies on the effects of glucagon, particularly in domesticated birds. Its role is thought to be well established:
• Stimulating lipolysis (chickens:
Langslow and Hales, 1969; turkeys:
Kurima et al., 1994; king penguins:
Bernard et al., 2003)
• Suppressing hepatic lipogenesis (chickens:
Goodridge, 1973; ducks:
Bedu et al., 2002).
• Elevating circulating concentrations of glucose (chickens: early research reviewed
Hazelwood, 1986; turkeys:
Kurima et al., 1994);
• Stimulating glycogenolysis in chick embryo hepatocytes (
Picardo and Dickson, 1982).
• Stimulating gluconeogenesis (discussed below).
In chickens and presumably other birds, the liver is the major organ converting lactate to glucose (
Watford, 1985). Glucagon stimulates hepatic gluconeogenesis (
Watford, 1985). On the other hand, the kidneys are the major gluconeogenic organ that uses either pyruvate or amino-acids as the substrate (
Watford, 1985). Renal cytosolic phosphoenol pyruvate carboxykinase activity is increased by cAMP (
Watford, 1989). What is not known is whether glucagon or other metabolic hormones act at physiologic concentrations to influence renal gluconeogenesis. For instance, does administration of glucagon antagonists suppress the elevated gluconeogenesis observed in the kidneys of fasted birds. Moreover, effects of glucagon antagonists on liver gluconeogenesis have not been reported.
Although glucagon can markedly elevate circulating concentrations of glucose in birds that does not per sedemonstrate that glucagon is physiologically controlling circulating concentrations of glucose. Glucagon does not appear to be playing a role in the tonic control of circulating concentrations of glucose at least in the fed state. Circulating concentrations of glucose were unaffected by administration of a glucagon antagonist in fed chickens (
Simon et al., 2000;
Dupont et al., 2008); this being when concentrations of glucagon are low (
Christensen et al., 2012a). What is not clear is whether blocking glucagon’s effect with an antagonist influences glycemia in the fasted state. Fasting markedly increases circulating concentrations of glucagon in chickens (
Christensen et al., 2012a). Moreover, it might be questioned whether glucagon when administered is mimicking other glucagon like peptides. There is further evidence that argue against glucagon being the only or even the principal hormone responsible for restoring homeostasis in the fasted state. The circulating concentrations of glucagon peak within six hours of fasting and decline as fasting is extended despite the increasingly severe relative hypoglycemia (Table 2). It is noted that the effects of glucagon on chicken adipose tissue are attenuated over prolonged exposure to glucagon (
Campbell and Scanes, 1987;
Oscar, 1992) due to downregulation of the glucagon receptors (
Oscar, 1996a).
The available but limited evidence of the role of glucagon in wild birds is consistent with its role in poultry. For instance, glucagon increases circulating concentrations of non-esterified fatty acids in red-winged blackbirds indicating stimulation of lipolysis (Hintz 2000). There is clearly a strong case for fundamental studies on the role of glucagon and other hormones in wild species of birds. It is argued that metabolism in poultry is markedly different from the ancestral wild state due to intensive selection.
The effects of glucagon on lipolysis are influenced by other hormones with pancreatic polypeptide and triiodothyronine (T3) being stimulatory while somatostatin (SRIF) and growth hormone (GH) are inhibitory (this is discussed in more detail under each hormone below).
The factors controlling secretion of glucagon in domesticated birds are summarized in Table 3. It is presumed that similar control mechanisms exist in wild species. For instance, glucose is reported to depress circulating concentrations of glucagon in migratory garden warblers (
Totzke et al., 1998). What is not known in birds are effects of gastric hormones such as the incretins-glucose dependent insulin tropic peptide (GIP) and glucagon like peptides (GLP) 1, together with gastrin and cholecystokinin, other gastro-intestinal peptides such as ghrelin, GLP-2, obestatin, and orexigen together with adipose hormones such as the ligand for the avian leptin receptor and adiponectin. Moreover, there is a need to investigate the physiologic roles of these controlling circulating concentrations following a meal or in fasting in birds using glucagon antagonists. What is furtherunknown in birds is the extent that the assays in the studies where peripheral glucagon was determined (
Laurent and Mialhe, 1978;
Honey et al., 1979) also detected other glucagon gene products such as GLP1 and 2, glicentin or oxyntomodulin.
Insulin
Insulin appears to lack several of the roles it has in mammals:
• No effect on lipolysis at least in domestic birds (based on
in vitro studies with chicken adipose explants (
Langslow and Hales, 1969),
• No effect on hepatic glycogenolysis based on studies in chick embryo hepatocytes (
Picardo and Dickson, 1982),
• No effect onglycolysis by either skeletal muscle or liver or gluconeogenesis by either liver or kidneys (
Chida et al., 2000).
However, insulin does have some effects in birds based on studies where either insulin was administered or
in vitro studies. Insulin administration increases muscle glycogen (
Hazelwood and Lorenz, 1959) despite the absence of the insulin dependent glucose transporter 4 (GLUT 4) in birds (chicken:
Kono et al., 2005; sparrow:
Sweazea and Braun, 2006a). Insulin enhances glucose entry into kidney, brain and muscle tissue but not adipose tissue in birds. This is supported by the decreased circulating concentrations of glucose following insulin administration (
Hazelwood and Lorenz,1959), the increased uptake of 2-deoxy-D-[1-(3)H]glucose ([(3)H]2DG) by skeletal muscles (
soleus, extensordigitorum longus and
pectoralis superficialis muscles)
in vivo (
Tokushima et al., 2005) and
in vitro (
Rudas and Scanes, 1983). Moreover, insulin does increase uptake of fatty acids by sparrow muscle (
Sweazea and Braun, 2006b).
Insulin increases expression of the liver form of phosphofructokinase in chicken breast muscle (
pectoralis) (
Seki et al., 2006). In addition, insulin
in vitro has a modest effect on the synthesis of fatty acids; insulin synergizing with T
3 to stimulate the synthesis of fatty acid synthase in the chicken liver (
Wilson et al., 1986) but has no discernible effect on lipogenesis by duck hepatocytes (
Bedu et al., 2002).
There is strong evidence for insulin having a physiologic role in the control of glucose metabolism in birds. For instance, circulating concentrations of glucose in fed chickens rise to extremely high levels when endogenous insulin is selectively removed using insulin antibodies (
Simon et al., 2000;
Dupont et al., 2008). Moreover, despite the elevated ambient glucose concentrations, there are very high circulating concentrations of glucagon (
Dupont et al., 2008). These data are consistent with insulin per se acting in a tonic manner to depress circulating concentrations of glucose. Alternatively, insulin antibodies can be envisioned tobe removing insulin’s tonic suppression of glucagon secretion(Table 3) which in turn elevates circulating concentrations of glucose. What is not clear is whether blocking glucagon’s effect with an antagonist can influence circulating concentrations of glucose in the hyperglycemic and hyperglucagonemic state following administration of antisera to insulin. Furthermore, it is not known the extent that insulin is mediating the increases in circulating of T
3andhepatic GH binding and the decreases in hepatic deiodinase type III activity following re-feeding fasted chickens (
Buyse et al., 2002). This could be approached by examining
in vivo using antisera to insulin.
Insulin also downregulates insulin receptor expression in the chicken hypothalamus and probably other tissues (
Shiraishi et al., 2011).Insulin influences release of insulin like growth factors; stimulating per se secretion of IGF-I by chicken hepatocytes
in vitro and acts synergistically with GH to increase IGF-I release (
Houston and O’Neill, 1991). Insulin also depresses hepatic IGF binding protein 2 expression in fasted chickens (
Nagao et al., 2001).
Table 4 summarizes the factors controlling insulin secretion in birds. What is not known in birds are effects of gastric hormones such as the incretins- GIP and GLP 1, together with gastrin and cholecystokinin, other gastro-intestinal peptides such as ghrelin, GLP-2, obestatin, and orexigen together with adipose hormones such as the ligand for the avian leptin receptor and adiponectin. Moreover, there is a need to investigate the physiologic roles of these controlling circulating concentrations of gastro-intestinal and pancreatic hormones following a meal or in fasting in birds using administration of specific antagonists orantisera.
In turkey poults, hypophysectomy decreases circulating concentrations of insulin with the effect partially overcome by GH replacement therapy (
Proudman et al., 1994).
Other pancreatic hormones
There are effects of other pancreatic hormones on metabolism. For instance, pancreatic polypeptide stimulates both basal and glucagon stimulated lipolysis by chicken adipocytes (
Oscar, 1993). There is scant information on the physiologic role for pancreatic polypeptide in birds. The metabolic effects of SRIF will be discussed under gastro-intestinal hormones (below).
Pituitary, thyroid hormones, adrenal hormones and metabolism
Glucocorticoid- corticosterone
Avian metabolism is profoundly influenced by either corticosterone, the major glucocorticoid in birds, or dexamethasone acting as a surrogate.Administration of glucocorticoids to chickens is followed by decreased growth, particularly of skeletal muscle with marked increases in adiposity and liver weight (
Bartov, 1985;
Siegel et al., 1989;
Jiang et al., 2008;
Yuan et al., 2008). Corticosterone decreases skeletal muscle and nitrogen retention in chickens (
Bartov, 1985). This is not apparently due to decreases in the rate of protein synthesis (
Hayashi et al., 1994). Rather, corticosterone is increasing degradation of both myofibrillar and non-myofibrillar muscle proteins (
Hayashi et al., 1994;
Ohtsuka et al., 1998;
Dong et al., 2007). Moreover, there are increases in the circulating concentrations of glucose, insulin, triglyceride and non-esterified fatty acids (NEFA) with glucocorticoid administration (
Yuan et al., 2008). There are also increases in adipose lipoprotein lipase (
Jiang et al., 2008) and
pectoralis muscle glycogen (
Lin et al., 2007;
Gao et al., 2008). Glucocorticoids decrease expression of IGF1 and increase that of myostatin in chicken muscle (
Song et al., 2011).It is unknown whether effects of the effects of glucocorticoid on avian metabolism are physiologic or do they represent supra-physiologic or pharmacological effects.
Glucocorticoids increase hepatic glycogen and circulating concentrations of glucose (
Kobayashi et al., 1989). There is evidence that glucocorticoids stimulate gluconeogenesis in the liver (
Watford, 1985;
Niwa et al., 1986) and kidneys of chickens (
Watford, 1989). There is clearly a need to examine whether or not physiologic concentrations of corticosterone stimulate gluconeogenesis in the liver (
Niwa et al., 1986) and kidneys of bird. This is particularly germane to carnivorous birds where gluconeogenesis would be anticipated to be more important. Moreover, it is be important to determine quantitatively the importance of liver and kidney to overall gluconeogenesis under different physiologic states and hormonal milieu.Glucocorticoids are thought to act by inducing insulin resistance in chickens (
Dupont et al., 1999;
Yuan et al., 2008). There are reductions in hepatic insulin binding and insulin receptor substrate-1 levels (
Dupont et al., 1999) together with decreased insulin stimulated glucose uptake in skeletal muscle (
Zhao et al., 2009a).
Growth hormone
Components of the hypothalamic pituitary growth hormone axis influence lipid metabolism. Ghrelin and SRIF are discussed under gastro-intestinal hormones (below). Lipolysis is increased by GH as indicated by elevated non-esterified fatty acids (hypophysectomized chickens:
Scanes et al., 1987; acutely in chickens:
Scanes, 1992; chronically in turkeys:
Bacon et al., 1995). Moreover, GH stimulates lipolysis
in vitro with chicken adipose explants (
Harvey et al., 1977;
Campbell and Scanes, 1985). GH from birds, mammals and reptiles is effective while that from lower vertebrates is not (
Campbell and Scanes, 1985;
Campbell et al., 1990;
1991). The effects of GH on lipolysis are blocked by a specific GH antagonist (
Campbell et al., 1993), by protein synthesis inhibitors and low calcium in the incubation media (
Campbell and Scanes, 1988a) and by either SRIF or insulin (
Campbell and Scanes, 1988b). In addition, GH has an anti-lipolytic effect with the lipolytic effect of glucagon
in vitro being reduced in the presence of GH (
Campbell and Scanes, 1987). The anti-lipolytic effect is observed with mammalian, avian GH together with both lower vertebrate GH (
Campbell et al., 1991) and the GH antagonist (
Campbell et al., 1993); the later not evoking a lipolytic effect. It is not known whether GH acts via direct effects on the adipocytes or by acting on fibroblasts or other cells found in adipose explants. In addition, GH depresses lipogenesis (
Harvey et al., 1977;
Rosebrough et al., 1991). There is a strong case to re-examine the effects of GH
in vitro determining effects on gene expression, protein synthesis, post-translational changes (particular phosphorylation) and employing a comprehensive metabolomics approach.
Growth hormone also influences metabolism indirectly by increasing circulating concentrations of IGF-1 (
Houston and O’Neill, 1991) and T
3 (
Darras et al., 1993).
Prolactin
Prolactin increases feeding and hence elevates the quantity of energy stored in adipose tissue in birds. This is supported by the ability of intra-cerebroventricular administration of prolactin to stimulate food consumption and increase bodyweight in ring doves (
Buntin et al., 1999). The effect is independent of glucocorticoids as prolactin induced hyperphagia is not blocked by a glucocorticoid antagonist (
Koch et al., 2004). It is not clear whether this is an effect of pituitary prolactinper se or of the recently described prolactin-like protein (PLP) (
Wang et al., 2010); PLP being expressed in the brain of chickens and also zebra fish (
Wang et al., 2010).
Thyroid hormones
The active thyroid hormone, T
3, has marked effects on metabolism in birds based on albeit relatively small number of studies in poultry species. Fasting results in decreases in body temperature (
Christensen et al., 2012b), presumably reflecting decreased metabolic rate, and reductions in circulating concentrations of T
3 (
Buyse et al., 2002). Re-feeding fasted chickens is followed by increased circulating concentrations of T
3 due to decreased hepatic deiodinase type III activity and elevated metabolic rate and particularly anaerobic metabolism (
Buyse et al., 2002). It is reasonable to propose that the increased T
3 is mediating the shift in metabolism. However, definitive studies are required to examine this hypothesis.
The transition from fasting to the fed state is accompanied by a shift from lipolysis to lipogenesis (
Buyse et al., 2002). It is unknown the extent to which shifts in the circulating concentrations of T
3mediate the changes in lipid metabolism. There is evidence that enhances T
3 both lipolysis and lipogenesis in birds. Pre incubation of chicken adipocytes with T
3 increases basal and glucagon stimulated lipolysis (
Suniga and Oscar, 1994). Lipogenesis in the chicken liver is stimulated by T
3 with increases in both the expression and synthesis of fatty acid synthase particularly in the presence of insulin (
Wilson et al., 1986).
Adrenocorticotropic hormone, β lipotropin and other products of proopiomelanocortin (POMC)
The avian pituitary gland expresses proopiomelanocortin (POMC), the precursor for ACTH,β-lipotropin, β endorphin and α, β and γ melanocyte stimulating hormone (MSH) (chicken:
Berghman et al., 1998;
Takeuchi et al., 1999; ostrich:
Naudé et al., 2006). The primary structure of ostrich β-lipotropin has been characterized (
Naudé et al., 1981). What is not known is the extent to which POMC products other than ACTH such as α, β and γ MSH and β-lipotropin are released from the avian pituitary gland. Both glucagon and ACTH stimulate lipolysis by chicken adipose tissue
in vitro (
Langslow and Hales, 1969;
Strosser et al., 1983). What is also not known is whether these hormones stimulate lipolysis in all avian species, whether the effects are physiologic and whether other products of the POMC gene such as α, β and γ MSH or β-lipotropin also influence lipolysis and metabolism in other peripheral organs.
Adipose hormones
Adiponectin
There is high expression of the adiponectin gene in chicken adipose tissue and to a less extent in the liver. Expression in both tissues is increased in chickens fasted for 48 h (
Maddineni et al., 2005). However, no changes in circulating concentrations of adiponectin were observed in fasted chickens (
Hendricks et al., 2009). An increase in adiponectin expression is reported in protein restricted chickens (
Tahmoorespur et al., 2010). In the chicken, there are two genes for the adiponectin receptor, namely AdipoR1 and AdipoR2. Both receptors are expressed in the adipose tissue together with skeletal muscle and liver with increased expression of AdipoR2 in adipose tissue in chickens fasted for 48 h (
Ramachandran et al., 2007). What is not known includes whether adiponectin has a physiologic role in either influencing carbohydrate or lipid metabolism directly or indirectly via effects on the secretion of insulin, glucagon or other hormones that control metabolism in birds.
Leptin
There are marked effects of leptin reported on metabolism in birds. For instance, administration of leptin increases hepatic expression of fatty acid synthase in chickens (
Dridi et al., 2005). The leptin receptor has been characterized (chicken:
Liu et al., 2007; goose:
Wang et al., 2011; turkey:
Richards and Poch, 2003). There is evidence for a physiologic role of the avian leptin receptors. Oleic acid increases expression of leptin receptors in goose adipocytes and expression of genes related to adipocyte differentiation are downregulated in adipocytes in which leptin receptor is knocked down (
Wang et al., 2011). There is high expression of the leptin receptor in the chicken brain (
Wang et al., 2011) with changes in hypothalamic expression during development (
Huang et al., 2010). The chicken leptin receptor activates the JAK-STAT pathway (
Adachi et al., 2008). The endogenous ligand for the leptin receptor is, perhaps surprisingly, not conclusively established in chickens or other species of birds(
Adachi et al., 2008). A bioassay for leptin failed to detect any leptin activity in chicken serum (
Hen et al., 2008). It has been argued that the chicken leptin structure deposited in Genebank is erroneous and that leptin has been lost in birds (
Sharp et al., 2008;
Pitel et al., 2010). Clarification is required as to the very existence of leptin in birds and to potential alternative ligands for the leptin receptor.
Gastro-intestinal Hormones
Ghrelin
There is evidence for effects of ghrelin on avian lipid metabolism. Ghrelin decreases hepatic expression of fatty acid synthase (
Buyse et al., 2009). Ghrelin presumably acting via the GH secretagogue receptor (GHSR) influences metabolism in birds. It is unknown whether other products of the ghrelin gene namely unacylated ghrelin or obestatin have physiologic roles in birds.
Somatostatin
There is evidence fordirect effects ofSRIF on metabolism in birds. Somatostatin suppresses glucagon stimulated lipolysis with chicken adipose tissue
in vitro (
Di Scala-Guenot et al., 1985;
Oscar, 1996b). In turkeys, SRIFhas been demonstrated to decrease glucagon induced lipolysis
in vivo (
Kurima et al., 1994). Similarly, SRIFinhibits GH stimulated lipolysis by chicken adipose tissue explants
in vitro (
Campbell and Scanes, 1988b). It is noted that SRIF is without any discernible effect on carbohydrate metabolism (
Kurima et al., 1994).What is not known whether avian lipolysis is physiologically influenced by SRIF. Moreover, it is questioned whether cortistatin which binds to SRIF receptors is also an anti-lipolytic hormone in birds.
Other intestinal hormones
It is not clear whether there are effects of gastro-intestinal peptides such as ghrelin, motilin or gastrin releasing peptide on avian lipolysis. It is further unknown whether growth factors influence avian lipolysis in an analogous manner to the effects of fibroblast growth factor 21 (FGF21) on human adipocytes (
Arner et al., 2008).
Control of carbohydrate metabolism in wild birds
Circulating concentrations of glucose and their control
The physiologic relevance of the comparative high plasma glucose concentrations exhibited by birds is not clear. The results of a number of studies have demonstrated these concentrations do not change significantly during or as result of long distance migration (
Jenni and Jenni-Eiermann, 1998;
Maillet and Weber, 2006) and that migrants “burn” primarily fatty acids. For example, Warblers migrating from Northern Europe to sub-Saharan Africa do so in “hop-Scottish” manner (
Schaub and Jenni, 2001).That is they tend to accumulate large fat depots before departing the summer feeding and breeding grounds, fly until the fat deposits are depleted, come to rest and replenish fat deposits (
Schaub and Jenni, 2001). This pattern is repeated several times until they reach their wintering territories. A second example is that of the Plovers (
Calidris pusilla) that are summer resident in Eastern Canada. These birds feed on the amphipod (
Corophium volutator) that contains high concentrations of n-3 polyunsaturated fatty acids. Once conditions are correct, they fly non-stop to New Zealand and their plasma glucose concentrations on arrival are not significantly different from that of departure levels (
Maillet and Weber, 2006). Thus, it remains somewhat of an open question as to why this condition has evolved in birds and how their tissues withstand plasma glucose concentrations that would be lethal to mammalsand to humans in particular.
The results of some interesting recent work demonstrated that small birds, Swainson’s thrushes (
Catharus ustulatu), flown in a wind tunnel at low relative humidity metabolized protein to produce water to prevent dehydration while sparing glucose (
Gerson and Guglielmo, 2011). These results add another intriguing dimension to the as yet not completely understood complex metabolism of birds.
Another interesting aspect of avian glucose homeostasis is that this group appears to lack Glut 4, the primary insulin responsive glucose transport protein on muscle (skeletal and cardiac) and adipose tissue (
Sweazea and Braun, 2006a). Moreover, birds appear to store very little glucose as glycogen in their tissues (
Ferguson and Braun, 2010).So why do birds maintain relatively high plasma glucose concentrations?
Metabolism related to flight and migration
The prevailing view is that high energetic needs or “fuel” for flight by birds are met predominantly by oxidation of fatty acids.
Short-term flights
The view that birds use fatty acids to “fuel” flight is supported by a relative small number of reports on short-term flights of less than 50 km. For instance, immediately following flight in homing pigeons, there are marked increases in the circulating levels of glucose, lactate, non-esterified fatty acids, epinephrine, glucagon and norepinephrine but no change in those of corticosterone and decreases in those of both T
3 and T
4 (
Viswanathan et al., 1987;
George et al., 1989). The view that flight is powered by fatty acids is supported by the observed respiratory quotient of 0.7 in fasted hummingbirds (
Suarez et al., 2011). However, glucose also can be the source of energy for flight as indicated by the respiratory quotient of 1.0 in nectar feeding hummingbirds (
Suarez et al., 2011). Moreover in pigeons, there is a high capacity for glycolysis in the type 1 red muscle fibers in the pectoralis flight muscles (
Meléndez-Morales et al., 2009).
Extended Flights in Migrations
The energetic needs for extended migrations are confounded by the absence of feeding during the period. Fatty acids are the major source of energy for flight during migration of birds (
Jenni and Jenni-Eiermann, 1998). This is not surprising given the energy density of triglyceride in adipose tissue:
• Glycogen in skeletal muscle and liver 3.5 - 4.4 MJ per gram wet weight
• Protein in skeletal muscle 5.3 MJ per gram wet weight
• Triglyceride in adipose tissue 37.6 MJ per gram wet weight
(calculated from Jenni and Jenni-Eiermann, 1998).
In migratory birds, there is a seasonal fattening prior to migration to provide stored energy for during the rigors of migration. Prior to migration, birds prepare for the anticipated energetic requirements by laying down adipose fat stores (e.g.
Landys et al., 2004). The hormonal basis for this is only partially understood. During and prior to migration, there are increases in circulating concentrations of the major avian glucocorticoid, corticosterone (
Landys et al., 2004;
Falsone et al., 2009).This is accompanying hyperphagia due at least in part to increased circulating concentrations of corticosterone based on studies with glucocorticoid antagonist (
Landys et al., 2004). There are shifts in the glucagon:insulin ratio(
Totzke et al., 2000). Circulating concentrations of glucagon are somewhat depressed during pre-migratory fat deposition in red-winged blackbirds (
Hintz, 2000). There are likely to be also reductions in circulating concentrations of T
3. Flight is associated with a reduction in basal metabolic rate in birds (
Speakman and Selman, 2003).
In addition in preparation for migration, there are changes in the expression of metabolic enzymes to prepare for the rigors of long distant flight. For instance, there is upregulation of both enzymes for fatty acid oxidation and the fatty acid transporters (membrane and intra-cellular) in flight muscles (e.g.
Bishop et al., 1995;
Pelsers et al., 1999;
Guglielmo et al., 2002;
McFarlan et al., 2009;
Guglielmo, 2010). Moreover, as might be expected, fatty acid synthase expression is decreased immediately prior to migration (
Zajac et al., 2011).
Birds use fatty acids as the major energy source for flight during migration (e.g. (
Sweazea and Braun, 2006a;
Vaillancourt and Weber, 2007). The corollary to this is that there should be the following:
• Adequate stores of triglyceride in adipose tissue,
• High rates of lipolysis and consequently elevated circulating concentrations of non-esterified fatty acids,
• High capacity for muscles to transport fatty acids and for β fatty acid oxidation
• Gluconeogenesis using amino-acids from protein degradation to supply glucose.
In migrating birds, there are very high rates of lipolysis (e.g.
Vaillancourt and Weber, 2007). Similarly at the end of migration of a shorebird, circulating concentrations of triglyceride, non-esterified fatty acids and glycerol are elevated; the latter two indicating high lipolytic activity (
Landys et al., 2005). Moreover, high circulating concentrations of uric acid supported protein degradation presumably to supply gluconeogenic precursors (
Landys et al., 2005).
Responses in extended periods of fasting
King penguins and other species of penguins will fast for periods of five months in the winter losing about two thirds of their bodyweight (
Cherel and Le Maho, 1985). There are three metabolic phases in response to prolonged fasting in birds:
• Phase 1: Adaptation with catabolism of fat stores,
• Phase II: Lower metabolism (economy) with slow but steady body mass loss using adipose lipid stores but with protein sparing,
• Phase III: More rapid bodyweight loss with elevated circulating uric acid indicating protein (muscle) degradation induced by the elevated circulating concentrations of corticosterone (
Spée et al., 2011).
During phase II, there are reductions in metabolic rate as indicated decreases in plasma
PCO
2 in studies in geese (
Le Maho et al., 1981). Moreover, plasma concentrations of the gluconeogenic amino-acid, alanine, are not changed (0.4 mM) (
Le Maho et al., 1981). This would indicate protein sparing. There are increases in lipolysis and fatty acid oxidation. Plasma concentrations of beta-hydroxybutyrate are considerably elevated by prolonged fasting in geese (
Le Maho et al., 1981) and king penguins (
Cherel and Le Maho, 1985). Perhaps surprisingly, glucagon is not maximally stimulating lipolysis and gluconeogenesis in fasted penguins as glucagon infusion increases lipolysis further (
Bernard et al., 2003). Moreover, glucagon infusion is accompanied by elevated circulating concentrations of glucose, beta-hydroxybutyrate and insulin in fasted king penguins (
Bernard et al., 2003).
During phase III of fasting, there are further shifts in the circulating concentrations of nutrients and metabolites. Prolonged fasting is accompanied by some decrease in the circulating concentrations of glucose and the gluconeogenic amino-acid, alanine(
Cherel and Le Maho, 1985).This is explicable by increases in protein (muscle) degradation generating alanine and other gluconeogenic amino-acids but a relatively larger increase in the rate of gluconeogenesis in the penguins. Similarly, circulating concentrations of glucose are maintained in the range 144-180 mg/dL (8-10 mM) in long-term fasted geese (
Le Maho et al., 1981). These changes would appear to be mediated induced by the elevated circulating concentrations of corticosterone (
Spée et al., 2011).
Control of metabolism in poultry
Plasma concentrations of glucose in poultry
There is a marked variation in the reported circulating concentrations of glucose in chickens even in the same physiologic state. For instance, in an analysis of 15 studies of plasma concentration of glucose in chickens of a limited age range with similar nutrition and genetics, mean plasma concentration of glucose were reported to vary between 156 and 330 mg/dL (
Scanes, 2008). In chick embryos, the disparity between studies/investigators is even more apparent with circulating concentrations of glucose being reported at 200 mg/dL on days 18 of embryonic development (
Lu et al., 2007) but at 111 mg/dL, when glucose was determined by isotope dilution gas chromatography mass spectrometry (
Sunny and Bequette, 2010). The basis for the disparities between reported glucose concentrations requires attention and the use of common standards. An issue that should be addressed is the capability of avian red blood cells to metabolize glucose. Unlike mammalian erythrocytes, avian red blood cells contain a nucleus and mitochondria. If avian erythrocytes utilize glucose for their energy needs, it is readily apparent that failure to separate red blood cells from plasma or to cool blood samples rapidly might result in erroneously low circulating concentrations of glucose.
Circulating concentrations of glucose rise during development of the chicken embryo; increasing from 120 mg/dL on days 10 to 12 to about 200 mg/dL on days 18 to 20 of embryonic development in the (
Lu et al., 2007). There were changes in circulating concentrations of glucagon but these temporally lagging those of glucose in chick embryo development (
Lu et al., 2007). Circulating concentrations of glucose increase markedly in the first 2 days of post hatching life in chickens rising from 140 mg/dl to 220 mg/dl with marked decrease in circulating concentrations of glucagon (
Richards et al., 2010). The circulating concentrations of glucose were reported to decline during growth in three populations of chickens (mean at 3 weeks old- 260 mg/dL; 6 weeks old -243 mg/dL; 9 weeks old -178 mg/dL; 12 weeks old -162 mg/dL (
Sinsigalli et al., 1987). Moreover, the incremental change in circulating concentrations of glucose declines 20 min after glucose load (2 g per kg intubated into the crop); the Δ glucose being at 3 weeks old-104 mg/dL; 6 weeks old -91 mg/dL; 9 weeks old -60 mg/dL; 12 weeks old -25 mg/dL (
Sinsigalli et al., 1987).
It is not clear as to what are the relative contributions of glucose, lactate and fatty acids to avian metabolism. Based on
in vivo studies where radioactive glucose was placed into a duodenal loop, about 37% of the glucose is converted to lactate by the wall of the small intestine during absorption (
Riesenfeld et al., 1982). Moreover, there is fermentation of starch to lactate in the crop of gallinaceous birds. This is illustrated in Table 5 for turkeys with similar levels of lactate in the chicken crop (
Moore et al., 2004).Circulating concentrations of lactate are reported at 43 mg/dL and muscle tissue levels of 3.6 mg/g (
Savenije et al., 2002). There is a need to determine the importance to overall avian metabolism of lactate either generated in the crop or small intestine wall. Moreover, if avian erythrocytes utilize glycolysis for their energy needs, it is readily apparent that failure to separate red blood cells from plasma or to cool blood samples rapidly might result in erroneously high circulating concentrations of lactate.
The role of liver and muscle glycogen in glucose metabolism
It is suggested that both liver and muscle glycogen play an important role supplying energy in birds during embryonic development and during the transition from
in ovo to independent life. During hatching of chicks, the concentration of glycogen in the liver declines (
Christensen et al., 2001). There are also decreases in the liver glycogen concentrations between days 15 and 19 of embryonic development (
Pulikanti et al., 2010). Glycogen in skeletal muscles declines during the process of hatching (
Christensen et al., 2001) presumably due to the energy needs for the physical process of hatching.
Glycogen is a readily available store of energy for times of feed interruption. There are reports that liver levels of glycogen decline during a fast. However, the available data exhibit a marked inconsistency in the rate of decline (>80% lost in 6 h -
Warriss et al., 1988; 77% in 24 h -
Edwards et al., 1999; 37% in 12 h -
Trampel et al., 2005). Moreover, levels of liver glycogen reported in the literature show marked variation between studies: 14 mg and 50 mg glycogen respectively prior to and following feeding in meal fed chickens (
Ekmay et al., 2010) and 1.2 and 2.6 g glycogen respectively prior to and following feeding in meal fed chickens (
de Beer et al., 2007). These differences may be explicable by the levels in the first study being a concentration (mg/g liver weight). There are decreases in muscle glycogen in chickens that had been subjected to feed withdrawal for 5 h; glycogen in the
pectoralisreported to decline from 15.5 to 8.6 µmoles per g (
Savenije et al., 2002).It is not clear whether this is µmoles glycogen or µmoles glucose generated by hydrolysis. Assuming that the glycogen concentrations reported are in glucose equivalents (
Savenije et al., 2002), then thenet rate of muscle glycogenolysis during fasting would be 246 µg per g per hour (Table 10) which is similar to the blood glucose concentration. There is a need for studies on glycogen levels, synthesis and degradation in the liver and skeletal muscle in fed and fasted adult birds and during growth. Moreover,
post mortem increases in muscle glycogen were reported (
Savenije et al., 2002). It is therefore essential to ensure tissues are rapidly frozen following sampling.
The role of liver fat in metabolism
The liver is a major site of fatty acid synthesis (Table 5). Hatching is accompanied by marked shifts in metabolism. With the transition from very high lipid ingested from the yolk to a starch based diet, there is a large increase in lipogenesis and in the expression of key enzymes in the lipogenic pathways (e.g. malic enzyme, fatty acid synthase and stearoyl coenzyme A desaturase) in the first few days of post hatching life in chickens (
Richards et al., 2010).
There are changes in liver fat and hepatic lipogenesis following feeding in adult chickens. The amount of fat in the liveris increased (from 3 g to 5.2 g) 12 h after feeding in adult female chickens fed daily (
Ekmay et al., 2010). A larger increase in liver fat (from 2.1 to 6.0 g)is reported in in birds fed on alternate days (
Ekmay et al., 2010). It is reasonable to suggest that these are due to lipogenesis from glucose and consequent triglyceride formation. Synthesis of fatty acids from
14C-acetate
in vivo is reported to be increased following feeding (Silpananta and Goodridge, 1971). Hepatic expression of both fatty acid synthase and malic enzyme are also increased after feeding (
Ekmay et al., 2010).
A model for metabolism
Table 5 summarizes metabolism in fed and fasted adult chickens. Fasting reduces whole body glucose utilization (
Belo et al., 1976) reflecting decreases in lipogenesis, triglyceride formation, muscle and hepatic glycogenesis and glucose oxidation. In meal fed chickens, there is net glycogen formation after a meal with much higher rate (170 mg/hour/kg bodyweight) in chickens fed on alternate days (
de Beer et al., 2007). Twelve hours following meal feeding, net glycogenolysis occurs with again a higher rate (250 mg/hours/kg). (
de Beer et al., 2007). Glycogenolysis is insufficient to maintain circulating concentrations of glucose during fasting. Given the estimates for glycogenolysis and glucose utilization, rates of gluconeogenesis are likely to be very high during fasting (Rate of gluconeogenesis= Whole body glucose utilization – Rate of Glycogenolysis).
Although there is not information on glucose oxidation by different organs, fatty acid synthesis
in vivo drops by 90% within 2 h of fasting (
Yeh and Leveille, 1970;
Goodridge et al., 1986). Decreases in glucose utilization by muscle tissue during fasting are supported by the reduction in arterial-venous glucose difference in glucose concentration in fasted 8 week old chickens (
Tinker et al., 1986). Similarly, there is a decrease in the metabolic rate in fasted growing chickens based on the reduced circulating bicarbonate concentration (Christensen et al., 2012b). Moreover although the rates of lipolysis or triglyceride formation are not established, circulating concentrations of free fatty acids were elevated within 30 min of fasting indicating increased lipolysis and decreased triglyceride formation (
Yeh and Leveille, 1970). It is noted that there is substantial storage of lipid in the liver (Table 5). During fasting, liver lipid is mobilized rapidly. On a weight basis, the rate of liver lipid breakdown is similar to that of whole body glycogenolysis (Table 5). However on an energetic basis, it would seem that lipid breakdown in the liver alone contributes considerable more than glycogenolysis.
Along with the limited information on metabolism in adult chickens, there is a surprising dearth of quantitative information on metabolism in young rapidly growing chickens. To model metabolism fully in birds, it will be necessary to quantify carbohydrate, lipids and protein. This should include the following:
• Characterizing carbohydrate metabolism, determining changes in amount of glycogen in the liver and muscles with time during fasting and following feeding together with rates of glycogenolysis, glycogenesis and glyconeogenesis throughout growth.
• Characterizing lipid metabolism, determining changes in amount of triglyceride in the liver and adipose tissue with time during fasting and following feeding together with rates of fatty acid synthesis, fatty acid oxidation, triglyceride formation and breakdown throughout growth.
• Characterization protein metabolism, determining changes in amount of protein in liver, muscles and gastro-intestinal tract with time during fasting and following feeding together with rates of protein synthesis and degradation throughout growth.
Responses to fasting
Feed deprivation can be addressed by a series of strategies:
• Mobilization of energy from chemical storage
o Glucose from glycogen
o Fatty acids together with glycerol from triglyceride
o Amino-acids from proteins as gluconeogenic precursors
• Reduced metabolic rate with potentially a consequent reduction in core body temperature
• Mechanic storage of feed including slower gut transit time
• Reduced locomotory activity and other behavioral changes.
The Role of mobilization of energy from chemical storage
The effects of fasting on metabolism are summarized in Table 5. Fasting has a relatively small effect on circulating concentrations of glucose in chickens perhaps explaining inconsistency between different studies. No effects of fasting on circulating concentrations of glucose have been reported in some studies (e.g.
Belo et al., 1976;
Brady et al., 1978). In contrast, decreases in circulating concentrations of glucose were in chickens were reported to be depressed by fasting (
Langslow et al., 1970;
Harvey et al., 1978;
Simon et al., 2011;
Christensen et al., 2012a) (see Table 2). Fasting in chickens is followed by decreases in both the concentration of glycogen in the liver (
Hazelwood and Lorenz, 1959) and in glucose replacement rate (
Belo et al., 1976). Moreover, there are increases in the circulating concentrations of free fatty acids (
Langslow et al., 1970). Fasting is associated with increases in circulating concentrations of amino-acids (
Belo et al., 1976;
Brady et al., 1978) reflecting protein degradation and providing a substrate for gluconeogenesis. There is evidence for increased gluconeogenesis in fasting in chickens with elevated kidney phosphoenolpyruvate carboxykinase (
Bisbis et al., 1994). Fasting reduces lipogenesis and increases lipolysis (
Buyse et al., 2002). In meat type chickens, fasting results in reductions in circulating concentrations of triglyceride and increases in circulating concentrations of fatty acids (
Nijdam et al., 2005;
Delezie et al., 2007). In an analogous manner, there is a large decline (>50%) in NEFA 20 min after start of meal (
Buyse et al., 2002).
Endocrine responses to fasting
In meat type chickens, fasting for as little as12 h results in marked reductions in circulating concentrations of insulin (
Shiraishi et al., 2011;
Simon et al., 2011;
Christensen et al., 2012a)(Table 2) and T
3 (
Buyse et al., 2002). Although circulating concentrations of insulin are depressed in fasted chickens, there are increases in insulin receptors in liver (
Simon et al., 1986) and kidney (
Bisbis et al., 1994). There appear to be no changes in circulating concentrations of corticosterone with fasting (
Nijdam et al., 2005;
Delezie et al., 2007). Fasting increases circulating concentrations of both glucagon (
Christensen et al., 2012a) and GH (
Harvey et al., 1978). Fasting increases hepatic IGF binding protein 2 expression in chickens (
Nagao et al., 2001). What is not precisely known are the roles of glucagon, insulin and T
3 in fasted poultry?
Metabolic rate and body temperature responses to fasting
Nocturnal period of fasting
In birds active during the day, core body temperatures peak during the day and are low at night (pigeons:
Graf et al., 1989; chickens:
Christensen et al., 2012b). In nocturnally active species, core body temperatures peak during the scotophase (rats:
Yoda et al. 2000). There is obvious selective value to reduce energy consumption during the period of inactivity, sleep and fasting. There are circadian changes in core body temperatures with lower temperatures during the period of inactivity (
Aschoff, 1983).The nocturnal fast is accompanied by reduced energy expenditures in humans (
Holmbäck et al., 2003) and chickens with the metabolic rate declining rapidly by 26% at night (
Buyse et al., 1993).
Short-term fasting
Fasted pigeons show reductions in both heat production and core body temperature with declines in both the zenith and nadir in the daily cycle of temperatures (
Graf et al., 1989;
Ostheim, 1992). Similarly in fasted chickens, there are marked decreased in core body temperatures (
Christensen et al., 2012b).Moreover, there is a nocturnal decrease in the metabolic rate in chickens with the rate at night being 26% lower than during the day (
Buyse et al., 1993).
Mechanical storage of energy
There is a unique capability for storage of feed in the crop and gizzard/proventriculus studied in gallinaceous birds such as chickens and turkeys; the crop being a blind sac in the lower esophagus; the gizzard being the equivalent of the muscular component of the mammalian stomach and the proventriculus being the equivalent of the glandular stomach. There is storage of largely or entirely undigested ingesta in the crop and also gizzard during the night when feeding activity is very low in chickens and turkeys (see Tables 6 and 7) (laying chickens:
Scanes et al., 1987; young broiler chickens:
Buyse et al., 1993; turkeys:
Cutler et al., 2005).The role of the gizzard in food storage is supported by the increased fill in both chickens and quail in which the crop has been surgically removed (
Savory, 1985).
Ingesta stored in the superior gastro-intestinal tract accounts for more than a half of the nocturnal energy needs of the poultry (
Scanes et al., 1987) (assuming metabolism is constant throughout the day and night):
• Crop storage accounting for 43% of the energy requirements of the laying chicken during the night,
• Storage of ingesta in the gizzard accounts for a further 14% of the energy requirements (
Scanes et al., 1987).
However, this does not take into account the nocturnal decrease in the metabolic rate in chickens (
Buyse et al., 1993). If we combine the decrease in metabolic rate and the feed/ingesta stored in the crop and gizzard, we can account for 77% of nocturnal energy needs.
Crop filling seems to depend on the amount of feed consumed.For instance, the crop is filled with feed in meal fed chickens and completely engorged in chickens meal fed on alternate days (
DeBeer et al., 2008). Moreover, the crop is similarly filled in meal fed turkeys (
Cutler et al., 2005;
Scanes, 2008). There is increased passage of feed into crop rather than the gizzard prior to the night in turkeys (
Jackson and Duke. 1995). The filled crop at the beginning of the period of night may be related to the high feed intake prior to lights off (
Savory, 1985). Surgical removal of the crop in Japanese quail and chickens influences feeding behavior with more frequent meals (
Savory, 1985). The rate of emptying of the crop (in g/hour) is similar in chickens meal fed daily or on alternate days (
DeBeer et al., 2008). This suggests an inherent mechanical process for voiding the crop of its contents. In addition, crop emptying can be influenced by nervous innervation, neuropeptides and dietary constituents:
• Ghrelin stimulates the muscles of the crop to contract and thereby the crop empty (
Kitazawa et al., 2009) while neuromedin C and bombesin, peripherally, depresses empting of the crop (
Tachibana et al., 2010).
• GLP-1, neuromedins B, C and bombesin, administered intracerebrally, reduces crop emptying (
Tachibana et al., 2003;
2010).
• Increased by tyrosine (
Furuse et al., 1991) or long-chain triacylglycerols (
Mabayo et al., 1992) but decreased by phenylalanine (
Furuse et al., 1991) and beta-hydroxybutyrate (
Furuse et al., 1997).
Nocturnal gut transit time represents another form of mechanical storage of feed. Indeed, gut transit time is reported to be markedly longer at night than during the day (
Buyse et al., 1993). It is not possible to different between this and crop/gizzard storage of ingesta. Gut transit time is closely related to filling and emptying of both the crop and the gizzard (
May et al., 1990,
1998).
Other responses to fasting
The fat mass and obesity-associated gene (FTO) has been characterized in poultry (
Jia et al., 2011). It is expressed in muscle, liver, adipose tissue, pituitary gland and the hypothalamus (
Jia et al., 2011). Fasting increases expression of the FTO together with that of the gene peroxisome proliferator-activated receptor γ coactivator 1α (PGC-1α)in muscle, liver and adipose tissue but deceases expression of both genes in the pituitary gland (
Jia et al., 2011). The role of the protein translated from transcript of the FTO gene in poultry and wild birds remains to be elucidated. Moreover, what is not clear whether all cells in the anterior pituitary gland express FTO transcripts and/or PGC-1α.
Overfeeding and metabolism
Ducks and geese are used for fatty liver as pate
foiegras. This is achieved by overfeeding. Overfeeding ducks with a high carbohydrate corn diet is accompanied by large, about 7 fold, in increases in liver weight in Pekin ducks, Muscovy ducks and hybrids (
Chartrin et al., 2006). There are increases in circulating concentrations of both glucose and insulin (
Berradi et al., 2007) and elevated hepatic specific activities of malic enzyme, acetyl coenzyme A carboxylase and glucose 6 phosphate dehydrogenase (
Chartrin et al., 2006). In addition overfeeding increases hepatic specific activity and expression of glucokinase in Muscovy and hybrids but not Pekin ducks (
Chartrin et al., 2006;
Berradi et al., 2007).
In ovo feeding
Increasing the availability of nutrients to the developing avian embryo influences both metabolism and post-hatching growth and development. In poultry, this is performed by inserting the nutrient solution into the amniotic sac and is referred to as
in ovo feeding. For example,
in ovo feeding with carbohydrate and β-hydroxy-β-methylbutyrate accelerates gastro-intestinal development and improves growth post-hatching in chickens(
Tako et al., 2004). Moreover,
in ovosupplementation with β-hydroxy-β-methylbutyrate increases both liver and muscle glycogen in turkey embryos (
Foye et al., 2006). Similarly,
in ovofeeding with dextrin and β-hydroxy-β-methylbutyrate increases hepatic glycogen, embryonic myoblast proliferation and post-hatch growth rate in chickens (
Kornasio et al., 2011)
Energetic needs for Immunity
It is assumed that there are energetic costs to both the innate and adaptive immunity with the costs greater when an animal is being challenged by the presence of a pathogenic or immune challenge (e.g.
Råberg et al., 2002). A corollary is that there will be trade-offs between the energetics needs for reproduction or growth andenergetics needs of immunity.
Is there an energetic cost for the existence of innate and adaptive immunity per se?
Surprisingly, there is evidence against for this conjecture at least for adaptive immunity. Transgenic mice lacking both B and T lymphocytes have a higher basal metabolic rate than controls in in a controlled but not germ-free environment (
Råberg et al., 2002). Råberg and colleagues (2002) concluded that “
energy saving is another plausible factor favoring the evolution of adaptive immunity.” In contrast, evidence for some energetic costs for the existence of innate and adaptive immunity include the protein synthetic and maintenance requirements of primary and secondary immune organs as these representing in aggregate about 1% of bodyweight (Table 8). What is not known is the energetic requirements in the face of an immune challenge.
What are the energetic costs for innate and adaptive immunity when challenged?
The available evidence generally supports there being increased energetic demands when immune challenged. For instance, the metabolic cost of the cell mediated immune response to phytohemogglutinin was estimated as 4.2 KJ per day or 29% of resting or basal metabolic rate in house sparrows (
Passer domesticus) (
Martin et al., 2006). In other passerine birds, the metabolic cost of the humoral immune responses (primary and secondary) was estimated as 8 - 13% of basal metabolic rate with a 2.8% decrease in body weight following challenge (
Swensson et al., 1998;
Ots et al., 2001) (see Table 8). In contrast, no increase in basal metabolic rate wasobserved in intact Japanese quail (
Coturnix coturnix) challenged with phytohemagglutinin (
Boughton et al., 2007). Basal metabolic rate tends to be further depressed in castrated quail following phytohemagglutinin challenge (
Boughton et al., 2007). Innate immunity will have significant he energetic demands. For instance, early host responses to an oral challenge with Salmonella Enteritidis include increases in macrophage number in the ceca and upregulation of expression of the inflammatory chemokines CXCLi1 and CXCLi2 (
Cheeseman et al., 2008a); these being ligands for the chicken CXCR1 (
Poh et al., 2008).. Chicken macrophages produce nitric oxide (NO) with increased inducible nitric oxide synthase expression in response to interferon-γ (
Cheeseman et al., 2008b).
Is there a required trade-off for growth due to the energetic costs for innate and adaptive immunity when challenged?
There is substantial evidence that a trade-off is required with growth due to the energetic costs for innate and adaptive immunity when challenged. First, decreases in growth correlate with increases in the constitutive innate humoral immune system in Leach's Storm-Petrel (
Oceanodroma leucorhoa) (
Mauck et al., 2005) and in tree sparrows (
Spizella arborea) (
Stambaugh et al., 2011).Secondly, chickens raised in a germ-free environment exhibit a markedly greater growth rate (23.4% calculated from review by
Lochmiller and Deerenberg, 2000). Thirdly, poultry selected for growth have depressed immune functioning. A meta-analysis has been conducted of immune-related responses in lines of chickens and turkeys selected for growth (
van der Most et al., 2011). This provides strong evidence that for a trade-off between utilizing energy for immune functioning and for growth. The meta-analysis excluded data from commercial lines, from “
articles presenting least-square means only and no raw means” and from
in vitro immune parameters. Moreover, mortality, antibody titers and a measure of cell mediated immunity were combined. In Table 9, combined data on mortality and morbidity from six of the studies in the meta-analysis are presented. Clearly, in both chickens and turkeys, there was greater mortality following infectious challenge in the high growth selected lines; by 3.4 fold in fast growing chickens (compared to a line selected for low growth) and by 1.94 fold in fast growing turkeys (compared to random bred line). Moreover, morbidity, as indicated by paralysis, was increased by 6.0 fold in the high growth rate chickens. Mortality and/or morbidity differences are presumed to reflect those of innate and adaptive immunity. It is not possible to preclude other factors such as tolerance of the disease organism(s) favoring the survival of the slower growing poultry.
Studies on differences in adaptive immunity between poultry selected for high growth and either random bred or slow growth line do not consistently support for the trade-off model. Data from a series of comparisons is summarized in Table 10. The weights of immune organs (thymus, bursa Fabricius and spleen) are greater in fast growing poultry but their relative weights tend to be lower. There are not consistently higher antibody titers in response to challenges. There are increased numbers of CD4
+/CD8
-peripheral blood lymphocytes in fast growing turkeys (
Li et al., 2000).
Another support for a trade-off between immunity and growth comes from the decrease in growth rate in chickens after a challenge to the adaptive immune system using a foreign protein (sheep red blood cells) or activation of the innate immune system in the acute inflammatory response to
Escherichia coli lipopolysaccharide (LPS) (
Klasing et al., 1987;
Mireles et al., 2005;
Zhang et al., 2011).
Are there trade-offs between the energetic needs for reproduction and for innate and adaptive immunity?
There is evidence supporting trade-offs between reproduction and immunity. There are marked seasonal changes both in reproduction and cell mediated immunity in house sparrows (
Passer domesticus) that are consistent with the view that there are trade-off between energy needs of reproduction and those of immunity with maximal immunity not being maintained throughout the year (
Martin et al., 2006). Moreover, castrated Japanese quail have a greater immune response than intact quail but a lower basal metabolic rate (
Boughton et al., 2007). Lipopolysaccharide (LPS) induces weight loss and reductions in feed intake in white-crowned sparrow (
Zonotrichia leucophrys) with the response greater in birds on long than short day lengths depending on body condition (
Owen-Ashley et al., 2008).
Are there trade-offs between the energetic requirements for responding to stressors and to the needs for innate and adaptive immunity?
There is abundant evidence that the avian glucocorticoid stress hormone, corticosterone, depresses immune functioning. In chickens, administration of corticosterone exerts the following effects:
• Decreased weight and weight relative to bodyweight of the bursa Febricius (
Davison et al., 1983;
1985;
Donker and Beuving, 1989).),
• Decreased weight and weight relative to bodyweight of the thymus (
Donker and Beuving, 1989) with increased apoptosis and expression of IL-1alpha, IL-6 and TNF-alpha (
Franchini et al., 2004),
• Reduced antibody formation (
Post et al., 2003),
• Antibody to infectious bronchitis virus (IBV) vaccination initially stimulated but thereafter titers decreased (
Shini et al., 2008),
• Depressed production of IgM while increasing that of IgG following challenge with West Nile virus (
Jankowski et al., 2010),
• Increased heterophil to lymphocyte (H/L) ratios (
Shini et al., 2008),
• Increased heterophils in circulation (
Post et al., 2003) but with reduced expression pro-inflammatory cytokines in heterophils (
Shini et al., 2010),
• Decreased number of lymphocytes in the circulation but with increased expression of interleukin (IL)-1beta, IL-6, IL-18, transforming growth factor (TGF)-beta4 and of chemokines CCLi2, CCL5, CCL16 and CXCLi1 in both peripheral and splenic lymphocytes (
Shini and Kaiser, 2009),
• Extended viremia following challenge with West Nile virus (
Jankowski et al., 2010),
In wild birds, corticosterone has similar effects. For instance, corticosterone reduces the antibody response but increases the innate immune response (red blood cell lysis) in owls (
Stier et al., 2009). Stress induced reduction in the immune response allows reallocation of resources to other critical physiologic functions.
Lipopolysaccharide (LPS) challenge
Lipopolysaccharide (LPS) challenge is accompanied after 6 h with initially hypothermia (core body temperatures reduced by 0.5°C) with reduced blood pressure followed by a rebound increase in body temperature (by 0.4°C compared to pre-challenge) (
Cheng et al., 2004;
De Boever et al., 2008,2009). Along with the LPS-induced inflammatory response in inducing fever, there is increased circulating concentrations of nitric oxide and corticosterone together with elevated expression of inflammatory response genes: TNF-like factor(TL)1A, Interleukin (IL)-1beta and IL-6 inducible NO synthase and toll-like receptor (TLR) 4 in the spleen and other lymphocytes (
Klasing et al., 1987;
De Boever et al., 2008,2009;
Takahashi et al., 2008). What is not known are the temporal changes in basal metabolic rate in birds challenged with LPS and the extent to which corticosterone (whose secretion is increased by LPS) reduces the inflammatory response to LPS.
Glutamine and immune function
Glutamine is utilized by rapidly proliferating immune and gastro-intestinal cells (
Calder and Yaqoob, 1999).It is not known the extent to which glutamine is utilized by avian lymphocytes following challenge or by the gastro-intestinal tract when undergoing regeneration following nutritional restriction and flight. Glutamine supplementation has little effect of the development of the small intestine during growth or following fasting in young chickens (
Yi et al., 2005).
Conclusions on the control of the control of circulating glucose in wild and domesticated birds including evolutionary aspects
When the plasma glucose concentrations of birds are compared to a closely related phylogenetic group of vertebrates, reptiles one might expect to observe similar plasma concentrations. Quite the opposite appears to be the case. In a data set compiled for 55 reptilian species by Dessauer (1970), the mean plasma glucose concentration was 80 mg/dL (Table 11), a value similar to that of mammals as phylogenetic group. Thus, the ability of birds to maintain and tolerate comparatively high plasma glucose concentrations appears to have evolved independent of other vertebrate groups.
Figure 2 summarizes the circulating concentration of glucose and the evolutionary relationship between birds, mammals and the major groups of living reptiles. Circulating concentrations of glucose in free living reptiles are increasingly found to be very low compared to mammals. Circulating glucose in caught sea-turtles was reported as 18 mg/dl (1 mM) with lactate being higher at 3.4 mM (
Lutz and Dunbar-Cooper, 1987). Circulating concentrations of glucose in birds are normally held within tight range about a set point. For instance in ducks, there is little difference between the plasma concentrations of glucose in fed or fasted birds (Table 2). However, they are elevated by 15% by overfeeding (
Berradi et al., 2007) but are increased, by 2 fold, 30 min after an intramuscular glucose challenge in ducks (
Applegate et al., 1999). There are increasing circulating concentrations of glucose with age in pigeons (
Prinzinger and Misovic, 2010).
The circulating concentrations of glucose were even lower in recent studies where plasma concentrations of glucose were determined in free living reptiles immediately after capture (Table 1). Not only are circulating concentrations of glucose high in birds, but they are only modestly decreased by insulin challenge in birds being, for instance, decreased by 19% doves (
Sweazea et al., 2006) and 25-30% in chickens (e.g.
Langslow et al., 1970;
Harvey et al., 1978;
Zhao et al., 2009a). In contrast in both mammals and the very limited studies in reptiles, exogenous insulin reduced circulating concentrations of glucose to extremely low levels. In alligators, administration of insulin (either bovine or alligator insulin at dose of 400 μg per kg) is followed by a steady decline over 12 h in circulating concentrations of glucose to less than 35 mg/dL (
Lance et al., 1993).
It might be speculated that the high circulating concentrations in birds are a requirement for the energetic requirements of flight. Militating against this intuitively obvious but probably wrong assumption are the following. The circulating concentrations of glucose for flightless birds, including ostriches and penguins are well within the avian range (Table 1). Moreover, in the other flying vertebrates, namely bats, circulating concentrations of glucose are within but to the low end of the mammalian range being 54 mg/dl (3.0 mM) in nectar-feeding bats (
Kelm et al., 2011) and from an average of 15 to over 80 mg/dl in different months in insectivorous bats depending on their physiology (
Roy and Krishna, 2011). Very high circulating concentrations of glucose (450 mg/dl) are reported in nectar-feeding bats after feeding with rapid decreases when the animals fly following eating (
Kelm et al., 2011).
Evolution of insulin
It is reasonable to propose that the structure of chicken insulin is that of the common ancestor of birds and at least of two major groups of reptiles (Fig. 3). Chicken insulin (
Perler et al., 1980) is identical to that from the ostrich (Evans et al., 1988) and species from 10 different orders of birds: (Spheniformes) - chickens (
Gallus gallus), turkeys (
Meleagris gallopavo) (order Galliformes); common pigeon (
Columbia livia) (order Columbiformes); emperor penguin (
Aptenodytes forsteri)(order Spheniformes); petrel (
Pagodroma nivea), fulmar (
Fulmarus glacialis) and booby (
Sula sula)(order Procellariformes); kittiwake (
Rissa tridactyla), murre (
Uria aalge)(order Charadriiformes); owl (
Strix aluca)(order Strigiformes); red billed tropicbird (
Phaethon aethereus)(order Pelecaniformes); heron (
Nycticoraxnycti)(order Ciconiiformes); ostrich (
struthiocamelus) (order Struthioniformes) and emu (
Dromiceius novaehollandiae) (
Simon et al., 2004). Moreover, insulins from other orders of birds (excluding the anseriformes - ducks and geese) have simple amino-acid residue substitutions compared to chicken insulin (Fig. 3). In contrast, duck and chicken insulin differ from each other by three amino-acid residues (Fig. 3)(reviewed:
Conlon, 2000; chicken:
Perler et al., 1980; duck:
Markussen and Sundby, 1973;
Chevalier et al., 1996;
Simon et al., 2004).
Perhaps surprisingly, insulin from one group of reptiles, the turtles, is identical to that of chickens, and the putative avian model (turtles:
Conlon and Hicks, 1990;
Cascone et al., 1991). Moreover, alligator insulin has only 3 differences in the amino-acid sequence (
Lance et al., 1984) while tortoise insulin differs from alligator insulin by 3 amino-acid residues (
Wang et al., 1999). In contrast, python insulin has 18 substitutions compared with alligator and 20 substitutions compared with chicken (
Conlon et al., 1997). Based on the identity of amino-acid sequences in both the A and B chains, it is suggested that that the turtle/chicken/ostrich insulin represents an ancestral form dating back to the divergence of the ancestors of birds, crocodilians and turtles (Order Testudines, Super order Chelonida). The divergence between the ancestors of turtles/crocodiles and birds is over 225 million years ago being estimated at 228 million years ago (MYA)(
Hedges and Poling, 1999), 247 MYA (
Sanders and Lee, 2007) or 257 MYA (
Shen et al., 2011). The common ancestors of birds/turtles and crocodiles diverged from that of snakes and lizards significantly earlier; estimated as 245 MYA (
Hedges and Poling, 1999), 282 MYA (
Sanders and Lee, 2007) or 285 MYA (
Shen et al., 2011). The three amino-acid substitutions in ducks compared to the presumed ancestral form of insulin occurred presumably after the divergence of the ancestors of ducks and chickens (Figs. 2 and 3).While, duck insulin has a lower biologic activity compared to that of the chicken or mammal in mammalian systems, it has identical affinities to either chicken or duck insulin receptors (
Constans et al., 1991;
Chevalier et al., 1996). The conservatism of insulin in birds and two branches of reptiles argues for its critical importance to the biology in these groups.
Overall conclusions
Not only are the circulating concentrations of glucose are much higher in birds than in either mammals or reptiles, but also there are maintained with tight homeostatic control. The evolutionary basis for these high glucose concentrations is unclear. It does not appear to be a requirement for flight. Indeed, the principal fuel for avian flight is fatty acids. Moreover, the circulating concentrations of glucose are high in flightless birds and are not elevated in flying mammals, the bats. Recent studies that report on the circulating concentrations of glucose in reptiles indicate that its concentration is very low in the wild state but very labile to perturbations such as handling during capture. Circulating concentrations of glucose are greater in mammals than reptiles (Figs. 1 and 3) although not to the same extent as in birds. It is speculated that the increases are related in some way to homeothermy with body temperatures in birds markedly higher than those in mammals.
There is a profound need for quantitative studies on metabolism, and metabolomics, in both poultry and wild birds. Avian metabolism can be markedly influenced by hormones in a manner not identical to that in mammals. There is, however, a need to establish the extent to which these effects on physiologic.
Higher Education Press and Springer-Verlag Berlin Heidelberg
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