Endocrine PancreasJoëlle DupontUnité de Physiologie de la Reproduction et des Comportements, Institut National de la Recherche Agronomique, 37380 Nouzilly, France

Nicole Rideau and Jean SimonUnité de Recherches Avicoles, Institut National de la Recherche Agronomique, 37380 Nouzilly, France

Chapter 27

Sturkie’s Avian Physiology. Copyright © 2015 Elsevier Inc. All rights reserved.

27.1 INTRODUCTION

The endocrine pancreas of the chicken (and of birds in general) displays several peculiarities (Hazelwood, 2000; Simon, 1989). Before discussing the current knowledge, we will recall the basics from these previous reviews. Most of these are certainly connected and interacting.

l Birds have high plasma uric acid and glucose levels, despite normal plasma insulin levels (see Simon et al., 2011, for chicken). Such high plasma glucose levels (2–2.15 g/L in the fasting state) would be harmful for humans.

l In chickens and ducks, high doses of exogenous insulin are required to induce hypoglycemia, and huge doses are not lethal. The chicken features high blood glucose and low insulin sensitivity and, as will become apparent in this chapter, the weak insulin release elicited by glucose in the isolated chicken pancreas is characteristic of type 2 diabetes in humans.

l Experimental diabetes models are not available for chick-ens. Pancreatectomies are rarely complete; plasma glu-cose regulation is, however, impaired during re-feeding. Diabetogenic drugs (alloxan or streptozotocin) are inef-ficient, even when glycemia is decreased before strepto-zotocin injection. Some isolated cases of diabetes have been described in pet birds, although not cited here.

l Glucagon exerts a potent hyperglycemic effect; it is the lipolytic hormone in birds. The glucagon-induced cyclic adenosine monophosphate (cAMP) increase is compara-tively small but long-lived; the feedback decrease of cAMP by fatty acids appears inefficient in chicken adi-pocytes. Insulin lacks an antilipolytic effect in chicken adipocytes. Several peptides have been shown to have an antilipolytic effect in vitro: APP (avian pancreatic poly-peptide), somatostatin, gastrin, and GLI (gut glucagon-like immunoactivity); however, their function in chicken physiology has not been fully quantified in vivo.

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l Chicken insulin is “hyperactive”, whereas duck insulin is “hypoactive”. Chicken insulin has accordingly been suggested as more potent at inducing hypoglycemia in chickens than sheep or bovine insulin (in Hazelwood, 2000). The extent of the difference may have been overes-timated: at high concentrations, a similar nadir (maximal hypoglycemic effect) should be achieved irrespective of insulin type (this is the case for insulin binding and phos-phorylation of artificial substrate in solubilized chicken liver or brain insulin receptors).

l The importance of the insulin–glucagon ratio in the con-trol of metabolism has been demonstrated in chickens and ducks. In vivo, this ratio is governed by opposite insulin–glucagon variations in response to nutritional sta-tus (fed versus fasting). Glucagon–insulin interactions are, however, more complex. Insulin immuno-neutralization in fed chickens (i.e., a nutritional status for high insu-lin demand, a protocol previously developed in ducks; Mirsky et al., 1964 in Dupont et al., 2008) evokes a rapid (less than 30 min) and large rise in plasma glucose levels. Surprisingly, injection of a glucagon antagonist (des-His1 [Glu9] glucagon) has no effect on plasma glucose levels in fed chickens (Dupont et al., 2008); a hypoglycemic effect might be expected. This strongly suggests that in normally fed chickens, the control of plasma glucose level relies more on insulin than on glucagon (in Dupont et al., 2008, 2012). The presence of glucagon is, however, essen-tial: in the pathological model of “hypoglycemia-spiking mortality syndrome”, where pancreatic glucagon content is extensively depleted following a virus infection, chickens do not eat and ultimately die from starvation in a profound hypoglycemic status (Davis and Vasilatos-Younken, 1995). To inhibit pancreatic glucagon release, glucose requires the presence of insulin (Dupont et al., 2008), which confirms previous results obtained in ducks and later on in mammals (see Miahle’s group references in Simon, 1989).

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l Chicken (and bird) metabolism is also peculiar in several ways (as discussed throughout this book). In short, gluconeogenesis is active in liver and kidneys but appears able to be regulated only in kidneys. Liver is the organ for de novo lipogenesis.

l Insulin exerts pleiotropic effects in chickens during embry-onic and posthatch development. The major typical effects on carbohydrate, lipid, and protein metabolism have been demonstrated. Insulin stimulates glucose and amino acid transport in various cells. The insulin effect on glucose transport is, however, limited in muscle and still doubtful in chicken adipocytes. Insulin stimulation of amino acid transport requires protein synthesis. Insulin greatly stimu-lates liver lipogenesis and expression of lipogenic enzymes. In contrast, insulin stimulation of lipogenesis remains mar-ginal, at best, in chicken adipocytes (lipogenesis is also intrinsically low in human adipocytes). Glucagon counter-acts insulin stimulation of lipogenesis in chicken liver.

l Finally, birds have a high body temperature (about 42 °C in chickens) and a high metabolic rate and anabolism. They are able to adapt to long catabolic states and to sur-vive long periods without food.

This chapter offers a synthetic view of current knowledge, focusing on major aspects of glucagon and insulin control for the sake of brevity. References have been kept as few as possible; early major original contributions (including ours) for chickens and birds are acknowledged and listed only by the first author’s name and date, but not referenced, when quoted in recent publications. For evident reasons, knowledge of the role of the endocrine pancreas is far more advanced in mammals; on several occasions, it was neces-sary to refer to it. Again, to save space, these data are listed in the text by the first author’s name and date. Readers will need to connect to PubMed to have access to full references.

27.2 PANCREAS EMBRYOGENESIS AND DEVELOPMENT

27.2.1 Morphology of Avian Pancreas (Figure 27.1)

In most birds, the pancreas is situated in the duodenal loop and consists of three lobes: the dorsal, ventral, and splenic lobes. The splenic lobe is contiguous with the rest of the pancreas, but in some birds it can be completely separated. A fourth lobe, sometimes confusingly called the “third lobe”, is a part of the ventral lobe and is distinguishable in galliform birds (which include chickens and quail).

27.2.2 Distribution of Avian Pancreatic Endocrine Cells between the Different Pancreatic Lobes

The endocrine pancreas contains clusters of cells called the islets of Langerhans, which are distributed throughout

PART | V Endocrine Theme

the exocrine parenchyma. These represent only 1–2% of the whole gland. In contrast to mammals, in which only one type of islets occurs, three types of islets are found in birds: the light, the dark, and the mixed islets. They differ in shape, size, and staining characteristics. The light islets (or B-islets) remain clear after classical staining procedures (Heidenhain’s iron hematoxylin); they are essentially com-posed of β- and δ-cells secreting insulin and somatostatin, respectively, and occasionally very few α-cells secreting glucagon. The dark islets (or A-islets) are composed essen-tially of α- and δ-cells secreting glucagon and somatosta-tin, respectively. They are larger in size than the B-islets. The mixed islets are composed of numerous β-cells and a few α- and δ-cells. Some wild bird species have mixed islets (randomly distributed) rather than light and dark islets (Steiner et al., 2010). A fourth type of endocrine cell is found in the pancreas: the F-cells. These cells synthesize and release APP; they occur as single cells or in groups dissemi-nated in the exocrine parenchyma, mainly in the dorsal and ventral lobes. Some F-cells can be found within the islets. Isolated δ-cells (as numerous as F-cells) are also found as dispersed extra-insular endocrine elements. The splenic lobe and, to a lesser extent, the third lobe (when present) notably contain more islet tissue per unit volume than do the dorsal and ventral lobes (Rideau, 1988).

The feature unique to the avian pancreas is the predomi-nance of the glucagon-producing cells over the insulin- producing cells in the ratio of approximately 2:1 (see Andrew, 1984 in Manakova and Titlbach, 2007). In addition, somatostatin-producing δ-cells are also more numerous in the avian pancreas than in that of mammals (Rideau, 1988). The existence, distribution, and relative frequency of vari-ous (neuro-)endocrine cells (biotin, serotonin, neuropeptide

FIGURE 27.1 Morphology of the avian pancreas. The anatomical relationship in the chicken of the four lobes of the pancreas (shaded) and neighboring organs (blood supply and liver not shown) viewed from the dorsal aspect. (1) cystic duct; (2) hepatic duct; (3) dorsal pancreatic duct; (4) duct of the third lobe; and (5) ventral pancreatic duct. After Mikami, S.I., Ono, K., 1962. Glucagon deficiency induced by extirpation of alpha islets of the fowl pancreas. Endocrinology 71, 464–473; The Endocrine Society.

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Y (NPY), and chromogranin) have been demonstrated in the pancreas of avian species (Ku et al., 2000; Lucini et al., 2000).

The islet tissue is richly vascularized, irrigated by the pancreaticoduodenal artery and drained by the pancreatico-duodenal vein. Innervation of the pancreatic islets of birds is not as apparent as it is in mammals. Electron microscopic studies have clearly shown that the main endocrine cellu-lar types of the pancreas of the domestic fowl are associ-ated with nerve terminals. The nerve supply to the islets is, however, less abundant than that to the acinar cells; it is also less marked to the A-islets than to the B-islets. In con-trast, δ-cells are richly and directly innervated in chickens (Rideau, 1988).

27.2.3 Development of Avian Pancreas

The pancreas is of endodermal origin; it is formed from the embryonic foregut. In birds it arises from three buds: a dor-sal bud and right and left ventro-lateral buds ( Rawdon, 1998) that appear at the stage of 22–31 somites (ss) (56–67 h) and begin to fuse at day 7 of embryonic development (Hamburger and Hamilton stage 30 (HH30)) (Matsuura et al., 2009).

The determination of endoderm to form dorsal and ven-tral bud derivatives occurs before formation of the buds. In mammals, the signaling pathways underlying the pancre-atic development process include the Hedgehog system, the homeobox gene Pdx1, and Notch signaling. The vast major-ity of neurogenin-3 (ngn3)-expressing cells are committed to the endocrine lineage (Rosenberg et al., 2010). Two tran-scription factors, Pax4 and Arx, have competing roles in the commitment of the first of the ngn3-positive cells to gen-erate either the α/pancreatic polypeptide (PP)-cell lineage or the β/δ-cell lineage. Subsequently, within cells that are committed to the β/δ-cell lineage, the persistence of pax4 expression seems to select for the β-cell lineage, whereas suppression of pax4 expression selects for the δ-cell lineage (Gittes, 2009).

Early in vitro studies by Rawdon using the dorsal pan-creatic bud of 5 day chick embryo established that pancre-atic mesenchyme may play an instructive role with respect to the early development of the chick pancreas. Results also suggested that the notochord plays an early role in induc-tion of the pancreas, although experimental evidence was lacking (Rawdon, 1998). Studies of the past decade con-firmed and extended Rawdon’s observations (Rawdon, 1998). Prepancreatic endoderm first receives pancreas-inducing signals from the underlying somatic mesoderm, inducing Pdx1 expression as early as 4 ss (i.e., in 1.5 day old embryos) (Katsumoto et al., 2009). The regulation of pancreatic endocrine development by Notch is also found in chicken: inhibition of Notch signaling results in increased endocrine differentiation, whereas activation of the Notch1 receptor blocks endocrine development and maintains the proliferation of pancreatic progenitor cells in the embryonic

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chicken pancreas (Ahnfelt-Ronne et al., 2007). Neurogenin 3, which is transiently expressed in pancreatic endocrine progenitors, is responsible for the activation of a transcrip-tion factor cascade that ultimately leads to the maturation of endocrine cells in chicken endoderm (Rosenberg et al., 2010). The origin of the ventral pancreatic precursor cells has been studied less extensively; they are located at sites in which Pdx1 expression is detected by in situ hybridization (Matsuura et al., 2009).

Organization, volume, and ratio among the different endocrine cell types vary throughout embryonic develop-ment and postnatal life. The dorsal bud is the major source of insulin, glucagon, and somatostatin cells, whereas the ventral buds generate PP cells (Rawdon, 1998). In the dorsal bud, glucagon immunoreactive cells appear first at 2.25–2.5 days of incubation, then insulin cells at 3–5 days and somatostatin cells at 3.25 days (Manakova and Titlbach, 2007). PP cells appear only later, at 7 days in the splenic lobe (Cowap, 1985). In contrast to the dorsal bud, insulin cells have not been identified in prospective ventral bud deriva-tives until 13 days of incubation (Manakova and Titlbach, 2007). Development of exocrine tissue is delayed in com-parison to that of endocrine tissue (Manakova and Titlbach, 2007).

27.3 INSULIN AND GLUCAGON PEPTIDES

27.3.1 Insulin and Pre-pro-insulin, Pro-insulin, and C-Peptides

The chicken insulin gene was cloned and sequenced as early as 1980 (Perler et al. in Simon et al., 2004); it is localized on GGA5 and exhibits the “ancestral” structure containing two introns: intron-1 interrupts the 5′ untranslated region, and intron-2 (about 3.5 kb, versus 141–786 bp in other ani-mal species; Steiner et al, 1985 in Simon et al., 2004) inter-rupts the coding sequence. It encodes a pre-prohormone (i.e., pre-proinsulin) that contains the signal peptide (rich in hydrophobic residues), the insulin B-chain, the connecting peptide (C-peptide), and the insulin A-chain. To date, there is no evidence for the existence of a second insulin gene in birds (in Simon et al., 2004).

Insulin has been purified and sequenced in chickens, turkeys, ostriches, geese, and Pekin and Muscovy ducks (in Chevalier et al., 1996). The insulin gene has been cloned and sequenced in other bird species, including humming-bird and 28 other species (Fan et al., 1993 in Simon et al., 2004). From deduced amino acid sequences, bird insulins appear highly conserved. As compared to human insulin, minor neutral changes on the B-chain concern few residues at both ends: B1, B2, B3, B20, B27, or B30 (out of 30 posi-tions). On the A-chain, changes concern positions A8–10 (out of 21 positions). In our study (Simon et al., 2004), some uncertainty remains for the last six amino acids of

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the A-chain because the reverse primer was targeting the corresponding codons and the stop codon. Changes on the A-chain (at A8–10 positions) between bird and human insu-lins are not neutral. This region is the most variable and is considered as the most immunogenic part. Accordingly, most antibodies prepared against mammalian insulins do not cross-react with chicken or duck insulin.

At the A8 position, a histidine residue is found in most bird species, as opposed to a glutamic acid residue in six species of the Anseriform order and a threonine in human insulin. The A8 position is now thought to be involved in the insulin-binding site to the insulin receptor (site 1; Sajid et al., 2009); the synthesis of human insulin analogs wear-ing different residues at the A8 position confirmed an early hypothesis, namely, the presence of histidine at A8 confers the properties of chicken insulin to such analogs (high bind-ing affinity and low dissociation rate; see Weiss et al., 2001, 2002; Wan et al., 2004 and references quoted in Chevalier et al., 1996). Chicken insulin is the only natural super ana-log thus far described (at least twice as potent as human or pig insulin in bioassays or binding assays in many species; see Kemmler et al., 1978; Schauder and Buck, 1969; Simon et al., 1974, 1977 in Chevalier et al., 1996).

A8 glutamic acid, as found in the Anseriform order, increases the thermodynamic stability of the molecule, which should make “duck-type” insulins even more potent than “chicken-type” insulins (Weiss, 2001). In fact, the presence of negatively charged residues such as glutamic or aspartic acid at this position decreases the affinity of ana-logs for the receptor (Weiss, 2001). Duck insulin accord-ingly exhibits low affinity in rat liver membranes (Chevalier et al., 1996) and low bioactivity in rat fat cells (Moody, per-sonal communication). The fact that chicken or Pekin and Muscovy duck insulin receptors (IRs) do not discriminate so extensively, compared to potencies observed with rat IR, may be specific to bird IRs (see Section 27.5.2). So, thus far, only two types of insulin have been found in 30 bird species: the hyperactive “chicken-type” insulin and the hypoactive “duck-type” insulin. From alignments of nonmammalian insulins (see references in Simon et al., 2004), A8 histidine was present early in the ancestor insulin molecule and dis-appeared in mammals (which possibly decreases the risk of hypoglycemic events and tumorigenicity). The appearance of A8 glutamic acid in anseriform birds and some snakes may have occurred independently in these species.

As in other species, bird C-peptides have been much less conserved; 14 changes out of 28 amino acid residues are found in 29 bird species (Simon et al., 2004). In other animal species, C-peptides are longer (31–38 amino acids; see Gross et al., 1989; Wahren et al., 2000 in Simon et al., 2004). Whether the shortness of bird C-peptide modifies the rate and yield of insulin synthesis is unknown. C-peptide is linked to B- and A-insulin chains through two dibasic residues at both ends (RR or KR, as in other species). In

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mammals, proteolytic cleavage and removal of these two pairs are performed by specific endopeptidases: PC3/PC1 and PC2 prohormone convertases and carboxypeptidase E. Such endoproteases have been detected in chicken embryos using heterologous antibodies (see Hernandez-Sanchez et al., 2002; Teshigawara et al., 2001 in Simon et al., 2004). Though predicted to be 28 residues long in all bird spe-cies, purified duck (Anas platyrhyncos) C-peptide is only 26 amino acids long (Marksen and Sundby, 1973 in Simon et al., 2004). Such extensive cleavages are also suggested in all the birds studied (with a possible exception in Passer in Simon et al., 2004). In mammalian species, at least two acidic amino acids surrounding a hydrophobic residue are present at the NH2-terminal end of C-peptide (e.g., EAED, EVE, and EAE); such a structure has been found to be criti-cal for a normal rate of conversion of proinsulin to insulin. A very similar structure is observed in bird species studied (DVE, DIE, or DAE; Simon et al., 2004). C-peptide has long been considered as devoid of physiological functions on its own (other than structuring the precursor during insulin bio-synthesis). C-peptide is now considered to control e-NOS, Na+, K+-ATPase, alpha-enolase, and several transcription factors (Wahren et al., 2012; Ishii et al., 2012). The central segment (ELGGGPGAG) or the COOH-terminal pentapep-tide fragment (EGSLQ) of human C-peptide mimics the action of the entire peptide in several assays. The central part is very different in bird species, and C-terminal frag-ments are acidic (QEEYE, HEEYQ, QEEFE, or PEEYE). The C-peptide may therefore have acquired specific bio-logical functions only in mammals, unless the still putative receptor co-evolved in bird species.

The length of available pre-proinsulin gene reading frames is similar in all bird species (187 nucleotides at the 5′ end and 134 nucleotides at the 3′ end; Simon et al., 2004). In contrast, the size of the PCR products varies greatly (from 2.7 kb in Passeriformes (Passer, Turdus, and Pica) to about 4.5 kb in Paleognathae species (Struthio, Rea, and Dromi-ceius)). Phylogenetic analyses supported the monophyletic origin of birds and identified the Paleognathae group as an isolated group at 100%, strongly suggesting this group is basal. After the branching of Paleognathae, a gallo-anserae was identified (at 63%), whereas all other Neognathae clus-tered at 99%. If we accept that Paleognathae, rather than Passeriformes, are basal to Neornithes, intron-2 must have evolved from a very long ancient form (likely close to that found in present day Paleognathae species) toward the shorter form found in Passeriformes.

In humans, some insulin gene polymorphisms are associated with diabetes. Some polymorphisms have been described in chicken (Bennett et al., 2006; Qiu et al., 2006) and in duck (one paper in Chinese: Kong et al., 2008) insulin genes. In the chicken, polymorphisms (mostly single nucle-otide polymorphisms), thus far identified, are located in 5′- and 3′-UTR or intron. In these pioneer studies, significant

Chapter | 27 Endocrine Pancreas

associations were found between body weight at hatching, 28 days, and 55 weeks and small intestine length (but not with abdominal fat weight) at 13 weeks of age. Further stud-ies conducted at other ages and involving other physiologi-cal parameters may reveal other associations.

27.3.2 Glucagon and Glucagon-like Peptides

27.3.2.1 Glucagon Structure and Physiological Effects

Glucagon (a member of the vasoactive intestinal polypeptide (VIP)–secretin family of peptide hormones) is a 29 amino acid peptide that is very similar across species. In mammals, glucagon is encoded by a single proglucagon gene, and there is a single mRNA that generates a single protein precursor that contains glucagon and two structurally related peptides at its carboxyl terminus: glucagon-like peptide 1 (GLP-1) and glucagon-like peptide 2 (GLP-2). The posttranslational proteolytic processing of this unique precursor is tissue spe-cific. In the pancreas, glucagon is one of the major peptides released by the α cells of A-islets in response to low glucose levels, whereas GLP-1 and GLP-2, but not glucagon, are produced in intestinal L cells and the central nervous system.

Chicken glucagon is also a 29 amino acid peptide that is derived from a single proglucagon gene, which expresses multiple mRNA transcripts with different coding potentials (Richards and McMurtry, 2008). In the case of the domestic fowl, the pre-proglucagon precursor is 151 amino residues long. Chicken glucagon differs from rat and human gluca-gon by only one amino acid (ser versus asn at position 28) and from duck glucagon by two amino acids (ser versus asn at position 28 and thr versus ser at position 16). In contrast to mammals, the chicken proglucagon gene expresses two classes of proglucagon mRNA transcripts (PGA and PGB): class A mRNA (PGA) codes for glucagon and GLP-1 and resembles lower vertebrate transcripts such as those found in fish, while class B mRNA (PGB) codes for glucagon, GLP-1, and GLP-2 and is more like mammalian transcripts. Pancreas and proventriculus displayed the highest expres-sion levels of class A and class B mRNA. Two prohormone convertases (PCs: PC2 and PC1/3) cleave the precursor into active peptides: PC2 mRNA is predominantly expressed in the pancreas and proventriculus, whereas PC1/3 mRNA is more highly expressed in the duodenum and brain. Fast-ing and re-feeding have no effects on proglucagon mRNA expression in various chicken tissues: pancreas, proventric-ulus, duodenum, or whole brain. This indicates that tran-scriptional regulation does not account for acute changes in plasma glucagon resulting from short-term changes in energy status in birds (Richards and McMurtry, 2008, 2009). The proglucagon gene is also expressed in various neurons of the avian retina (Fischer et al., 2006) and in chicken adipose tissue (Cogburn, personal communication),

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whereas, in contrast to in other tissues, PGB mRNA levels appear insulin dependent (Ji et al., 2012).

In birds, glucagon has several physiological roles and peculiarities that already have been described in Hazelwood, 2000, 5th Edition. Plasma glucagon levels are much higher in birds than in mammals. Chicken, ostrich, and pigeon glu-cagons were initially found to be less potent than mamma-lian glucagon, but this was not confirmed in a subsequent study (see the following: McCumbee and Hazelwood, 1978; Ferreira et al., 1991; Tung et al., 1977; Huang et al., 1987 in Simon, 1995). In turkeys, pig and chicken glucagon were found to be equally effective in promoting hyperglycemia, an increase in plasma nonesterified free fatty acid levels, and a decline in circulating levels of T3 and T4 (McMurtry et al., 1996). Long-term injection of glucagon has been shown to induce uncoupling protein (UCP) expression in the skeletal muscle of ducks, consistent with the thermogenic action of this hormone in birds (Raimbault et al., 2001). Glucagon is also important in the chicken eye for regulating ocular growth (Vessey et al., 2005). Honda et al. found that central, but not peripheral, administration of glucagon suppressed food intake and induced hyperglycemia in chickens and that these effects were mediated by the downstream actions of corticotrophin-releasing factor (CRF; Honda et al., 2007 in Honda et al., 2012). Thus, glucagon may serve as a neu-rotransmitter in the avian central nervous system, as has been suggested for mammals (Mayo et al., 2003).

27.3.2.2 GLP-1

GLP-1 slows down nutrient assimilation by inhibiting gas-tric emptying and acid secretion. Central but not peripheral administration of GLP-1 inhibits food intake and crop emp-tying in chickens and shifts fuel utilization from carbohy-drates to lipids without affecting overall energy expenditure (Tachibana et al., 2007). The hypothalamus is involved in the anorexic effect of GLP-1 in chicks (Tachibana et al., 2004 in Tachibana et al., 2007). Moreover, the anorexigenic effect of central GLP-1 administration is mediated by the GLP-1 receptor because co-administration of a specific GLP-1 receptor antagonist (exendin 5–39) or N-terminal fragments attenuates the anorexia, while injection of the antagonist alone increases food intake in layer-type but not broiler chicks (Furuse et al., 1998). These results indicate the potential for variable responses in food intake to endog-enous GLP-1 among different strains of poultry. Expres-sion of the GLP-1 receptor mRNA in chicken pancreas is consistent with a possible incretin role for GLP-1 in birds (although it is not yet demonstrated in chicken).

27.3.2.3 GLP-2

In mammals, GLP-2 plays a role in intestinal growth and nutrient absorption by maintaining the integrity of muco-sal epithelial cells. Secreted by enteroendocrine L-cells in

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response to the presence of nutrients, GLP-2 promotes crypt cell proliferation and suppresses apoptosis in mucosal epi-thelial cells. Elevated expression of GLP-2 receptor mRNA in chicken duodenum is consistent with the proposed role for GLP-2 in intestinal growth and function. Shousha et al. reported no effect of intracerebroventricular or intraperito-neal administration of GLP-2 on food intake, body tempera-ture, or locomotor activity in Japanese quail (Shousha et al., 2007). However, these studies involved the use of rat GLP-2, which shares only 52% amino acid identity with chicken GLP-2 and may lack activity in avian species.

27.4 INSULIN AND GLUCAGON RELEASE

27.4.1 Pancreatic Hormone Levels During Development and After Hatching

Circulating concentrations of insulin and glucagon rise dur-ing development of the chicken embryo. In Cobb 500 chick embryos and hatched chicks, plasma insulin increased from 10 days of embryonic life (E10) (30 pg/mL), reach-ing 1000 pg/mL at 17 days after hatching (Lu et al., 2007). Plasma insulin concentrations, at 3–4 ng/mL in fed grow-ing ducks and chickens, are similar to those of mammals (Simon et al., 2011). They are low at one day post hatch and increase with age up to 28 days, with higher insulin sensitivity in one-day-old chicks than in 21 day old broiler chickens, as observed in the latest strain of broiler chick-ens (Shiraishi et al., 2011b) and in previous reports on egg-laying or meat-type chickens. Plasma insulin concentration decreased by 45–50% after 6 h of fasting in 19 to 47 day old male and female modern meat-type chickens (Christensen et al., 2013). A low plasma glucagon concentration (59 pg/mL) was found in embryos at E10, remaining low until E17 and then increasing approximately threefold by the time of hatching (Lu et al., 2007). Values between 152 pg/mL and 450 pg/mL were measured in fed and 6 h fasted modern meat-type chickens, respectively (Christensen et al., 2013). It should be noted that in some of these experiments, the feeding pattern of chickens was not always synchronized with light–dark cycles.

27.4.2 Insulin Release

Whereas in mammals glucose is the primary physiological regulator of insulin secretion, the insulinotropic effect of glucose is less obvious in birds in vivo and in vitro. In vivo, raising blood glucose levels in ducks and chicken only tran-siently increases insulin levels (Rideau, 1988). The high basal plasma glucose concentrations found in chickens may themselves contribute to the relative insensitivity of the chicken β-cell to glucose: impairment of glucose-induced insulin release has been reported in hyperglycemic mam-malian models. The mechanisms are, however, presently

PART | V Endocrine Theme

unknown in chickens. Somatostatin, which is present at high concentration in the chicken pancreas (Weir, 1976 in Rideau, 1988), may also exert inhibitory paracrine effects on the β-cell and partly explain the insensitivity of the β-cell to fuel nutrients. However, somatostatin immunoneutraliza-tion in isolated-perfused chicken pancreas has only transi-tory effects on insulin release at high glucose concentration (42 mM) and no effect at low levels (14 mM) (Honey, 1981 in Rideau, 1988). In vitro studies provided further insight into the mechanisms responsible for the relative glucose insensitivity of chicken β-cell. Using isolated and perfused duodenum-pancreas from adult hens (King et al., 1976 in Rideau, 1988, 1998) first showed that the pancreatic β-cell is relatively insensitive to glucose as very high and nonphysi-ological glucose concentrations (30–40 mM) are required to induce a modest insulin release. This insensitivity was confirmed and extended to other fuel nutrients, which are potent insulinotropic agents in mammals, using isolated duodenum-pancreas from young chickens. d-glucose and d-glyceraldehyde require high concentrations to evoke either very weak or delayed insulin release; leucine and its ketoacid (α-KIC) are even totally ineffective. The chicken β-cell, however, remains sensitive to depolarizing agents (K+), tolbutamide (which closes K+ATP-dependent chan-nels), and potentiators of insulin secretion (acetylcholine or cAMP; Rideau, 1988). Therefore, the initiation of insu-lin secretion by fuel nutrients, the critical and specific step of the β-cell, appears different in chickens and mammals, whereas potentiating mechanisms (arginine, acetylcholine, and cAMP in the presence of low glucose concentration) and membrane depolarization events (K+ and arginine) are present in chickens as in mammals.

The key role of glucokinase (GK) in the regulation of insulin release and hepatic glucose utilization has been well described in mammals. Although the GK gene is not refer-enced in the chicken genome at Ensembl, we isolated a chicken GK cDNA and showed that the GK messenger is expressed in chicken liver and pancreas (Berradi, 2005 in Rideau et al., 2010). A human GK antibody identified a GK protein in chicken liver, the amount and activity of which were insulin dependent (Dupont et al., 2008). Furthermore, RO0281675 (a specific GK activator in mammals) caused severe hypogly-cemia in chickens but, surprisingly, did so without signifi-cantly increasing insulin levels, contrary to results obtained in mammals (Rideau et al., 2010). Thus, liver glucose utiliza-tion is substantially enhanced following liver GK activation. Although GK is present in the chicken pancreas, pancreatic GK activation may not be sufficient to cause insulin secretion, which confirms the in vitro studies reported here and supports the conclusion that the coupling between intra-B-islet metab-olism and insulin release is different in chicken B-islets. The lack of insulinotropic effect of leucine or α-KIC (metabolized in the mitochondria) suggests that the inefficiency is located beyond the pyruvate step in the chicken β-cell.

Chapter | 27 Endocrine Pancreas

Isolated chicken B-islets are necessary to further under-stand the insensitivity of the chicken β-cell to “insuli-notropic” fuel nutrients. Although functional islets were isolated in various mammalian species more than 40 years ago, isolation of avian islets of Langerhans has proven to be especially difficult due to the fact that the morphology of the isolated A- and B-islets had not been characterized. Ruffier et al. described isolation of functional A- (glucagon) chicken islets (Ruffier et al., 1998). Glucose-stimulated insulin release from B-islets isolated from the dorsal lobe of the chick pancreas has been reported (Datar et al., 2006). In our opinion, however, the specificity of these prepara-tions remains to be established as the dithiocarbazone stain-ing method stains both B- and A-islets in chickens (Ruffier et al., 1998).

Regarding resistance to diabetogenic drugs, the extreme resistance of birds to the diabetogenic drugs alloxan and streptozotocin (STZ) was described very early. In mammals, the β-cell specificity of these drugs is mainly due to the fact that they are selectively transported by the β-cell glucose transporter, GLUT2 (Lenzen, 2008). GLUT2 expression was reported in the liver and kidney in chickens; unfortu-nately, the pancreas was not examined (Kono et al., 2005). Within the mammalian β-cell, alloxan selectively inhibits glucose-induced insulin secretion through specific inhibi-tion of glucokinase; in addition, it induces ROS formation, resulting in the selective necrosis of β-cells. STZ’s effects are attributed to its specific alkylating potency, which modi-fies biological macromolecules and DNA fragments and ultimately destroys the β-cells (Lenzen, 2008). Compared with mouse islets, chicken islets generate smaller amounts of ROS under STZ. Consequently, less damage to proteins, lipids, and DNA can be expected in chicken than in mouse islets. These results may partly explain why STZ does not induce diabetes in chicken, even at high doses (Modak et al., 2007).

Despite the peculiarities present in chickens in the coupling between metabolism and insulin release, insulin release is under the control of multiple components in vivo and in vitro. Although glucose or a mixture of amino acids separately increases plasma insulin to a small extent, they act synergistically when combined in vivo. Re-feeding with gradually increasing amounts of food results in gradual rises in plasma insulin levels, adjusting glucose to a similar level. Changes in the glucose–insulin balance are observed in several experimental conditions or models (Table 27.1), including genetically fat and lean lines of chickens, dwarf chickens (dw/dw), hypothyroidism, corticosterone treat-ment, and chronic exposure to high temperature (Rideau, 1997). The incretin effect is not documented in chickens, but a variety of other stimuli, including nutrients (glucose and arginine), hormones (cholecystokinin, glucagon, corti-costerone, and GH) (Rideau, 1988), and neuronal stimuli (Karmann et al., 1992), increase plasma insulin levels.

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The effects of ghrelin–obestatin and GLP-2 on insulin release are not known in birds. Conversely, insulin secre-tion is inhibited by epinephrine and somatostatin (Rideau, 1988). Although the existence of leptin in the chicken is still debated, a recombinant “chicken” leptin (or leptin analog) potently inhibits insulin secretion from the perfused chicken pancreas (Benomar et al., 2003).

27.4.3 Glucagon Release

Fasting markedly increases circulating concentrations of glucagon in chickens (Edwards, 1999 in Scanes, 2009). In modern meat-type chickens, plasma glucagon levels increased 3.5- to 3.7-fold after 6 h of fasting while plasma glucose was depressed by 11% (Christensen et al., 2013). Glucagon secretion is inhibited by glucose in vivo in domes-tic birds (Honey, 1980 in Rideau, 1988) and in wild spe-cies (Tzotze, 1998 in Scanes and Braun, 2012). Exogenous glucose (42 mM) inhibited glucagon release by 27% from isolated A-islets of Langerhans (Ruffier et al., 1998). Con-versely, glucagon release is stimulated by amino acids, free fatty acids, cholecystokinin, somatostatin, and insulin in vivo (Scanes and Braun, 2012). A tonic suppression effect on glucagon by insulin is suggested in a model of insulin immune-neutralization in chickens (Dupont et al., 2008). As for insulin, the effect of gastrointestinal peptides and adipose tissue hormones on glucagon secretion is unknown in birds.

The mechanisms underlying the secretion of glucagon from α-cells are largely unknown in chickens. In mam-mals, α-cells contain a comparable secretory “machinery” as in β-cells: a glucose transporter (in this case, the ubiq-uitous GLUT-1), glucokinase, K+ATP channels, L-type voltage-gated calcium channels, and secretory granules (Le Marchand and Piston, 2010). However, α-cell secretory activity is quite different from that of β-cells. The mecha-nisms underlying the glucose inhibition of α-cell secretory activity are poorly understood. Two nonexclusive models have been proposed: a direct inhibition by glucose, and an intra-islet paracrine inhibition arising from non-α-cells. The paracrine control might involve somatostatin released from δ-cells and insulin, zinc, or gamma-aminobutyric acid released from β-cells. Recently, Le Marchand and Piston suggested that in the absence of cell–cell contacts, isolated α-cells function similarly to β-cells (i.e., glucagon secretion increases as a function of glucose concentration). However, in the islet, glucose inhibits glucagon release downstream the calcium step, presumably at the level of vesicle trafficking or exocytotic machinery (Le Marchand and Piston, 2010).

27.4.4 Somatostatin

Somatostatin (also known as growth hormone–inhibiting hormone (GHIH) or somatotropin release-inhibiting factor

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TABLE 27.1 Experimental or Genetical Models Exhibiting Changes in the Plasma Glucose–Insulin Relationship in Chickens1

Models

Fasting Plasma Levels Glucose Tolerance

Insulin Sensitivity FatteningGlucose Insulin Glucose Disposal Insulin Levels

Intermittent (IT) versus ad libitum feeding (C)

IT ≥ C IT = C IT > C IT < C ND IT ≥ C

Corticosterone (cort) treatment.

Cort > C Cort > C Cort < C Cort > C Cort < C Cort > C

Fat–lean lines (FL-LL) FL < LL FL ≥ LL FL > LL FL > LL FL ≥ LL FL > LL

High- and low-glucose lines (LGl–HGl)

LGl < HGl LGl = HGl LGl = HGl LGl = HGl LGl = HGl LGlL > HGlL

High- and low-growth lines (HG–LH)

HG ≤ LG HG > LG HG = LG HG > LG ND HG > LG

Dwarf (dw)–Normal (N) chickens

dw = N dw = N dw = N dw < N dw > N dw > N

1ND: not determined. Refeeding experiments have been performed following a fast in several of these models; results, not summarized in the table, can be found in Simon (1988) and in Simon et al. (2000) (LGl–HGl). Corticosterone was injected in doses of 5–6 mg/kg. Insulin sensitivity was evaluated through the hypoglycemic effect of exogenous insulin. Results of glucose tolerance in HG–LG chickens are unpublished data (Simon et al.).

(SRIF)) is a 14 amino acid peptide that regulates the endo-crine system. Chicken somatostatin is present in two forms, somatostatin-14 (SST-14) and somatostatin-28 (SST-28), which are encoded by the same gene (PSS1) as in mam-mals. Chicken pancreatic A-islets, B-islets, and D-cells contain only the SST-14 peptide form (Takayagani et al., 1996 in Trabucchi et al., 2003). Another somatostatin gene variant (PSS2) exists in many species, including chickens. PSS2 messenger is expressed in chicken pancreatic islets or pancreas and in a few specific brain structures, fewer than the structures expressing PSS1 messenger (Trabucchi et al., 2003).

27.4.5 Avian Pancreatic Polypeptide

APP was first isolated and characterized from chicken pan-creas (Kimmel et al., 1968); it is a member of the neuropep-tide Y family (PP, peptide YY (PYY), and NPY). PYY and PP are gut endocrine peptides almost exclusively expressed in the digestive system, whereas NPY is expressed in the central and peripheral nervous systems at all levels of the gut–brain axis. All three peptides are 36 amino acids long and act on G protein–coupled receptors; five receptor sub-types have been described. In mammals, PP is released from the endocrine pancreas following a meal and acts preferen-tially via Y4 receptors. It inhibits gastric emptying, intesti-nal electrolyte, water secretion intestinal motor activity, and peristalsis. PP also reduces appetite (Holzer et al., 2012). In chickens, APP is released in response to gut peptides and amino acids (Colca, 1982 in Rideau, 1988). Y4 receptors are expressed in a large number of peripheral tissues and in the brain (Lundell et al., 2002). APP primarily acts by

inhibiting gastrointestinal tract motility and gall bladder and exocrine pancreas secretion in chickens (Hazelwood, 1993 in Hazelwood, 2000). It also exerts metabolic effects: liver glycogenolysis, hypoglycerolemia, and lowering of the plasma free fatty acid level. One study, however, sug-gests that APP increases food intake in white leghorn chick-ens (Denbow et al., 1988).

27.5 GLUCAGON AND INSULIN RECEPTORS

27.5.1 Glucagon Receptors

In mammals, the biological actions of glucagon are medi-ated by a specific glucagon receptor (GCGR; Authier and Desbuquois, 2008), a receptor belonging to G protein–coupled receptor subfamily B, which also includes the receptors for GLP-1, GLP-2, VIP, growth hormone-releasing hormone, and growth hormone–releasing hormone-like peptide. Stimulation of the glucagon receptor modulates adenylate cyclase, initiates the production of cAMP, and thereby activates extracellular signal-regulated protein kinase-1 and -2 via cAMP-dependent protein kinase A (cAMP–PKA; Figure 27.2). In addition, it also activates signaling pathways via cAMP-independent interactions, leading to stimulation of phospholipase C and release of Ca2+ from IP3-sensitive intracellular Ca2+ stores.

The chicken glucagon receptor gene was cloned from adult brain tissue (Wang et al., 2008a); it encodes for a short glucagon receptor of 496 AA (GCGR-s) or a long receptor variant of 554 AA, designated glucagon receptor variant 1 (GCGR-v1). The chicken GCGR-s shares high AA sequence identity with that of human (70%), rat (69%), mouse (69%),

621Chapter | 27 Endocrine Pancreas

FIGURE 27.2 Glucagon receptor signaling.

and Xenopus (64%). Functional analysis confirmed that both GCGR-s and GCGR-v1 can be activated by glucagon and are functionally coupled to the cAMP-PKA signaling pathway. The glucagon receptor is widely expressed in all tissues, including the hypothalamus, with the greatest expression found in the liver, suggesting that glucagon plays a wide range of roles in both hepatic and nonhepatic chicken tissues through the two receptors, GCGR-s and GCGR-v1 (Richards and McMurtry, 2008). Recently, Wang et al. iden-tified a novel ligand–receptor pair, named GCGL and GCGL receptor (GCGLR), from chicken brain (Wang et al., 2012). GCGL is a novel GCG-like peptide of 29 amino acids that shares high amino acid sequence similarity with mammalian and chicken GCG (62–66%). GCGLR is a GCGL-specific receptor, which shares relatively high amino acid sequence identity with chicken GCGR and structurally related recep-tors (GLP1R, GIPR, and GLP2R). GCGL mRNA expres-sion is mainly detected in the central nervous system, spinal cord, and testis, whereas GCGLR is more widely expressed in chicken tissues (Wang et al., 2012).

27.5.2 Insulin Receptor

Early studies on ontogeny, structure, binding affinity, bind-ing characteristics, number, internalization, and up- and downregulation of insulin receptors (IR) in chicken or turkey tissues have already been reviewed (Simon, 1989).

Since then, changes in IR number have been described in the hypothalamus, at least at the level of IR messenger, in layer and broiler-type chicks under ad libitum conditions (Shiraishi et al., 2011a); changes in hypothalamic IR mRNA in response to photoperiodism are dependent upon gonad hormones (Anraku et al., 2007). In chicken liver plasma membranes, hepatocytes, and thymocytes, insulin-binding capacity (IR number) is lower than in rat or veal (in Simon, 1989), which may contribute to the low insulin sensitivity of chickens. However, in crude liver membranes (i.e., plasma and intracellular organite membranes), the IR number is about the same in chickens and rats (Dupont et al., 2004). A high proportion of IR may therefore be internalized in chicken liver (Krup and Lane, 1981, 1982 in Simon, 1989). Chicken IR shows a heterotetrameric structure (two α and two β subunits linked by disulfide bonds, as described in mammals and other species; reviewed in Simon, 1989). The α subunit (about 135 kDa) is entirely within the extracel-lular compartment and binds insulin; the β subunit (about 95 kDa) contains a large hydrophobic fragment (inserted into the membrane), crosses the cell membrane, and extends into the intracellular compartment. In chickens, as in other species, the α subunit of brain or muscle receptors is smaller than that of liver receptor α subunit, in part as a result of dif-ferences in glycosylation (reviewed in Simon, 1989).

In mammals, both IR α and β chains are synthesized from a single mRNA, which is encoded by 22 exons. In

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mammals, exon 11 is alternatively spliced, which results in two IR isoforms (IR-A and IR-B) that differ by the absence or presence of 12 amino acids at the C-terminus of the α subunit, respectively. IR-A and IR-B isoforms display dif-ferences in ligand affinity binding, kinase activity, receptor internalization, and recycling as well as intracellular sig-naling capacity and tissue distribution (De Meyts, 2012). According to Hernández-Sanchez et al., the region corre-sponding to exon-11 is missing in chicken IR; chicken tis-sues would only express the IR-A type (i.e., the IR type showing the highest insulin affinity) (Hernandez-Sanchez et al., 2008). This may account for the fact that chicken IRs exhibit much higher affinity for IGFs than mammalian IRs. It may also explain why chicken and duck IRs do not dis-criminate among chicken, pork, and duck insulins very well (Chevalier et al., 1996).

In its intracellular part, the IR β subunit is a tyrosine kinase activated by autophosphorylation following insulin binding to the α subunit. The chicken IR tyrosine kinase activity shows the same features as those from other species (reviewed in Kato et al., 2000; Simon, 1989). Chicken liver or kidney IR tyrosine kinase activities (basal and maximal activities) are regulated, for instance, by nutritional condi-tions (e.g., fasting or re-feeding), as in mammals. In con-trast, muscle and adipose tissue IR kinase activity, as well as β subunit tyrosine phosphorylation, do not decrease in the fasting state or increase in re-fed animals, despite altera-tions in plasma insulin (Adamo et al., 1987; Dupont et al., 2008, 2012). Thus, for still unknown reasons, the regulation of the tyrosine phosphorylation of IR β subunit and kinase activity differs between species and tissues. As depicted in Figure 27.3 (steps highlighted in blue for chicken tissues), once activated, the IR tyrosine kinase phosphorylates sev-eral intracellular substrates on specific tyrosine residues, which represents the proximal and early events initiating and propagating the insulin signal through a cascade of reactions leading to the multiple actions of insulin. Phos-phorylated IR substrates activate directly or indirectly various protein (serine–threonine) or lipid kinases, includ-ing phosphatidylinositol 3′-kinase (PI3K), protein kinase B (Akt), P70S6K, and mitogen-activated protein kinase (MAPK)–extracellular signal-regulated kinase (ERK)-1/2. Several of these postreceptor steps also regulate insulin cell sensitivity, enabling alterations of insulin sensitivity with-out changes in insulin receptor number or kinase activity.

27.5.2.1 Proximal IR Substrates

In mammals, at least 11 intracellular IR substrates have been identified (reviewed in Taniguchi et al., 2006; Siddle, 2012). These include the family of IRS (insulin receptor substrate) proteins (IRS-1–6) and Shc (Src homology col-lagen) protein. Four different IRS proteins (IRS-1–4) have been reported in various mammals, and two additional IRS

PART | V Endocrine Theme

proteins have been detected in humans (IRS5/DOK4 and IRS6/DOK5). These last two additional IRS proteins are involved in insulin signaling but are not involved in PI3K activation (the major insulin-signaling step). Other IR sub-strates such as Gab1, Dok, APS, SH2B, Cbl, and Crk have been also described in mammalian tissues or cells (cited by Siddle, 2012).

The coding region of chicken IRS-1 gene has been cloned and sequenced (Taouis et al., 1996). The deduced chicken IRS-1 protein (cIRS-1) presents high sequence identity with its human, rat, and mouse homologs. cIRS-1 is at least present in the liver, muscle, and adipose tissue and interacts with the insulin receptor (Dupont et al., 1998 in Dupont et al., 2009, 2012). Interestingly or surprisingly enough, the tyrosine phosphorylation of cIRS-1 and PI3K activity (as discussed further in this chapter) is accordingly dependent upon nutritional conditions or insulin status in the liver but not in muscle or adipose tissue (Dupont et al., 1998 in Dupont et al., 2009, 2008, 2012), suggesting a tissue-specific regulation and a relative insulin refrac-toriness in chicken muscle and adipose tissue. Coding sequences have recently been described for IRS-2, IRS-4, Crk, Cbl, and APS chicken homologs (Ensemble release 71, April 2013, at http://www.ensembl.org/Gallus_gallus/index.html); they are located on chromosomes 1, 4, 19, 1, and 19, respectively. However, the presence of these sub-strates as proteins and their insulin sensitivity remain to be established in chicken tissues. Cbl and Crk, which in mam-mals mediate insulin signals and are associated with lipid rafts, are upregulated, at the messenger level, in chicken adipose tissue by fasting (Ji et al., 2012). In mammals, this pathway stimulates glucose uptake, in addition to and independently of PI3K activation. Whether these poten-tial substrates contribute to the apparent refractoriness of PI3K activity to insulin that is observed in chicken skeletal muscle and adipose tissue remain unknown. It should be noticed, however, that insulin stimulation of glucose trans-port in chicken adipose tissue, if any, is very poor (as discussed further in this chapter).

In a chicken hepatoma cell line (LMH cells), cIRS-1 repression substantially increases Shc expression (Taouis et al., 1998). Shc is phosphorylated on tyrosine residues in response to insulin and therefore serves as an additional IR substrate in the absence of IRS-1. Chicken liver, muscle, and adipose tissue express homologs of the three mamma-lian Shc isoforms (Dupont et al., 2008, 2012). Interestingly, tyrosine phosphorylation of Shc (52 kDa) is dependent on insulin status: it increases after re-feeding or insulin injec-tion and is inhibited after insulin immuno-neutralization in chicken liver but also, surprisingly, in leg muscles, in con-trast to IRS-1. Also surprisingly, neither insulin privation nor fasting alters Shc tyrosine phosphorylation in adipose tissue (Dupont et al., 2009, 2012). In chicken liver, Shc, mainly the 52 kDa isoform, interacts with IR, whereas the

623Chapter | 27 Endocrine Pancreas

FIGURE 27.3 Insulin receptor signaling. (Adapted from mammals; only steps highlighted in blue have been characterized in chicken tissues).

46 kDa isoform associates with IRS-1 (Dupont et al., 1998 in Dupont et al., 2009). The 52 kDa Shc isoform also inter-acts with the p85 regulatory subunit of PI3K. Shc (52 kDa) is thus a key substrate for the insulin receptor in the chicken because, first, it is able to associate with PI3K and, sec-ond, in contrast to IRS-1, the extent of its tyrosine phos-phorylation is dependent on insulin levels. Furthermore, in liver, Shc (52 kDa) phosphorylation is inhibited after insulin immuno-neutralization (Dupont et al., 2008). The possible existence of different subtypes of IR complexes signaling through IRS-1 or Shc has been investigated in the liver. The presence of a large complex involving IR, IRS-1, Shc (mainly the 52 kDa isoform), and the regulatory subunits of PI3K has been demonstrated; and a third complex, includ-ing IRS-1 and the 46 kDa Shc isoform, is also present in chicken liver (Dupont et al., 1998 in Dupont et al., 2009). No such interactions have been described in mammalian tissues, and the functions of these complexes in chicken liver and their presence in other chicken tissues remain to be established.

27.5.2.2 PI3K–Akt

PI3K has been characterized in chicken liver, leg muscle, and adipose tissue (Dupont et al., 1998 in Dupont et al., 2009, 2012) and in LMH cells (Taouis et al., 1998 in

Dupont et al., 2009). Liver PI3K activity depends upon the nutritional state (prolonged fasting versus refeeding) in several experimental models (Dupont et al., 1998, 1999 in Dupont et al., 2009). Furthermore, PI3K activity increases in response to insulin injection and is inhibited follow-ing insulin immuno-neutralization (Dupont et al., 1998 in Dupont et al., 2009; see also Dupont et al., 2008). This is in agreement with the changes observed for early steps of the insulin-signaling cascade (tyrosine phosphorylation of IR β subunit, IRS-1, and Shc). Conversely, in muscle, the PI3K activity and, as previously mentioned, tyrosine phosphory-lation of IR and IRS-1 are totally unresponsive to insulin levels in experimental models that alter insulin sensitivity, such as long-term corticosterone treatment and genetically selected fat and lean lines of chickens (Dupont et al., 1999 in Dupont et al., 2009). As mentioned in this chapter, only the phosphorylation of the 52 kDa Shc isoform is reduced in muscle by fasting and increased by re-feeding. Compari-son of the responses to exogenous insulin in chickens and rats suggests that this phenomenon is specific to chicken. Furthermore, PI3K activity in leg muscle is about 30 times greater in the chicken than in the rat (Dupont et al., 2004). This “constitutive” hyperactivity of PI3K in chicken mus-cle may overstimulate the feedback inhibitory pathway described in mammals (as discussed in this chapter), thereby desensitizing chicken muscle to insulin. The p85 regulatory

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subunit of PI3K itself exerts strong inhibitory control of IRS signaling in mammals (Taniguchi et al., 2006). However, lipid phosphatases such as PTEN (phosphatase and tensin homolog deleted on chromosome 10) may also contribute to this relative insulin insensitivity of the early steps of insu-lin signaling in chicken muscle. We have detected PTEN in chicken tissues, including muscle (Vaudin et al., 2006); however, the role of PTEN in the control of insulin signal-ing in chicken tissues remains to be determined. In adipose tissue, insulin signaling is even more peculiar; insulin pri-vation did not alter early steps and PI3K activity, but also other downstream elements such Akt (see next paragraph; Dupont et al., 2012).

Akt has been detected in several chicken tissues includ-ing liver, muscle (Bigot et al., 2003; Dupont et al., 2008), adipose tissue, and granulosa cells (Dupont et al., 2012; Tosca et al., 2006). The phosphorylation of Akt on Ser 473 is increased following feeding of posthatched chicks (Bigot et al., 2003); this is prevented when feeding is delayed (48 h), then rapidly restored following food intake. In fed chickens, insulin immuno-neutralization significantly decreases Akt phosphorylation in both liver and muscle, but not in adipose tissue (Dupont et al., 2008, 2012). Thus, insulin is presumably involved in Akt activation in chicken muscle, despite the fact that chickens exhibit particular fea-tures in the early steps of the insulin-signaling cascade (IR, IRS-1, and PI3K). This issue remains unresolved. Poten-tial mechanisms involved in the atypical feature found for insulin signaling in chicken muscle and adipose tissue have been discussed in Dupont et al. (2008, 2012).

27.5.2.3 P70S6K

p70S6 kinase, a further downstream step, has been char-acterized in a quail muscle (QM7) cell line and in chicken skeletal muscle (Tesseraud et al., 2003, http://www.science direct.com/science/article/pii/S001664800800378X; Duchêne et al., 2008 in Dupont et al., 2009). p70S6K activity increases significantly in both pectoralis and gas-trocnemius muscles after re-feeding for 30 min following 16 h starvation, and also in fasting chickens after a single insulin injection (Bigot et al., 2003). Similarly, p70S6K activity in the muscle of neonatal chicks is correlated with the plasma insulin concentration, strongly suggesting an insulin-dependent p70S6K activation (Bigot et al., 2003). In the liver, stimulation of phosphorylation of P70S6K1 and activation of S6K (using specific artificial substrate) was not detectable (Dupont et al., 2008), although insulin stimulation of phosphorylation of S6 ribosomal protein was observed in the same conditions (Duchene et al., 2008b). Furthermore, p70S6K phosphorylation decreases in leg muscles but not in adipose tissue in response to insulin immuno-neutralization (Dupont et al., 2008, 2012). These findings and those for Akt are again surprising in view of

PART | V Endocrine Theme

the relative insulin refractoriness of chicken muscle to exog-enous insulin (Dupont et al., 2004). However, p70S6K has been found to be involved in another feedback loop, which attenuates the ability of insulin to activate PI3K. Indeed, there is evidence in mammals that p70S6K inhibits insulin signaling by increasing the serine–threonine phosphoryla-tion of IRS-1. In chicken muscle, re-feeding and insulin injection increase the phosphorylation of both p70S6K on key residues (T389, T229, and T421/S424) and its downstream target, the ribosomal protein S6 (downstream insulin-signaling steps). However, in these conditions, the phosphorylation of IRS1 on serine 632 and serine 635 resi-dues is also induced (Duchene et al., 2008b). In mammals, the phosphorylation of these sites inhibits the insulin signal. Thus, the AKT/p70S6K may inhibit IRS1 tyrosine phos-phorylation in chicken muscle.

27.5.2.4 MAPK–ERK1/2 Pathway

MAPK–ERK has been identified in various chicken tissues (Duchene et al., 2008a). In contrast to mammals, ERK1 is not present in bird tissues; ERK2 is the only phosphorylated isoform detected (Duchene et al., 2008a). It is activated in chicken myoblasts and LMH cells in vitro by insulin (Duchene et al., 2008a). U0126, an inhibitor of the MAPK–ERK pathway, completely abolishes insulin-induced S6K1 phosphorylation and activity in chicken myoblasts in vitro, implicating MAPK–ERK2 in the control of S6K1 by insu-lin in avian cells (Duchene et al., 2008a). MAPK–ERK2 phosphorylation in chicken liver muscle and adipose tis-sue in vivo is strongly reduced by insulin immuno-neutral-ization (Dupont et al., 2012, 2008). However, the role of MAPK–ERK2 in insulin action in vivo in chicken has not been investigated. This signaling could be crucial to chicken growth, and this pathway may participate in the insulin-dependent regulation of p70S6K, because Shc, which is upstream, seems to be a key IR substrate in chicken muscle.

From the elements of insulin signaling thus far studied in chickens, insulin signaling appears to work in chicken liver as expected from the knowledge gained from mam-mals. In contrast, in muscle and adipose tissue, the insu-lin cascade appears refractory, suggesting the presence of inhibitory counterregulatory mechanisms in these tissues.

27.6 GENERAL EFFECTS OF GLUCAGON AND INSULIN

27.6.1 Insulin and Embryonic or Posthatch Development

The insulin gene is expressed early during embryonic devel-opment in extra-pancreatic tissues prior to development of the pancreas, showing specific regulations (de la Rosa and de Pablo, 2011). Insulin accelerates the growth of embryos

Chapter | 27 Endocrine Pancreas

explanted at Hamburger Hamilton stage 4, without increas-ing mitoses; some specific developmental genes are upregu-lated (Patwardhan et al., 2004). A fine tuning of insulin gene expression is required to sustain adequate morphogenesis. In chicken neurulating embryos and chicken embryo retina, proinsulin is expressed but not cleaved to insulin. Whereas this precursor is considered to exert marginal (if any) bio-logical activity in peripheral tissues, it exerts anti-apoptotic effects in those tissues at this developmental stage, probably through a hybrid insulin–IGF receptor (in de la Rosa and de Pablo, 2011). Furthermore, two transcription-induced chimerisms have been identified with the adjacent tyrosine hydroxylase gene (TH-insulin; in de la Rosa and de Pablo, 2011). (In humans, a similar mechanism has been described between the insulin gene and IGF2, the other adjacent gene.) The insulin gene is also expressed in the neural tube and somites of chicken embryos. The addition of insulin (or IGFs) to other specific factors in the incubation medium cocktail stimulates somite cell proliferation and myo-genesis (Pirskanen et al., 2000). This effect is blocked by an insulin antibody and insulin receptor antibody, show-ing that insulin acts as a myogenic signal early in embryo-genesis. Insulin also supports in vitro multiplication of 1 day old broiler muscle satellite cells and delays their dif-ferentiation (Sato et al., 2012). Surprisingly, an opposite effect is observed between insulin and IGF1 for MyoD and myogenin messengers. Insulin or tolbutamide administra-tion to day-old broiler chicken increases Pax7 messenger (a striated cell marker) in breast muscle at 3 days of age and their body weight at 50 days without changing their body composition (Sato et al., 2012). These interesting effects deserve further characterization. In satellite cell–derived myotubes, insulin (at high doses, though, compared with IGF1) stimulates glucose (as discussed in this chapter) and amino acid transport and protein synthesis and inhibits pro-teolysis (Duclos et al., 1993a,b).

27.6.2 Insulin–Glucagon and Food Intake

Pioneer studies suggested that insulin contributes to the regulation of food intake (through inhibitory or stimula-tory mechanisms; in Simon, 1989). Intracerebroventricular injection of insulin (through the melanocortin system) or glucagon inhibits food intake in young leghorn chickens (Honda et al., 2007; Shiraishi et al., 2011b and quoted ref-erences). Surprisingly, insulin is inefficient in young broiler chickens in fasted or fed conditions (Shiraishi et al., 2011b and quoted references), suggesting a difference in their set point or sensitivity; in broiler chickens, plasma insulin lev-els are higher, but they appear to decrease hypothalamic IR mRNA expression (Shiraishi et al., 2011b). Insulin immuno-neutralization in fed growing chickens decreased food intake, most likely by inducing very high plasma glu-cose levels (Dupont et al., 2008).

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27.6.3 Insulin and the Endocrine System

Insulin interacts with glucagon (see Section 27.1) and with other endocrine systems. As early as 1 h after insulin immuno-neutralization (Dupont et al., 2008), plasma T3 levels decreased (no reliable measurements were obtained for T4). After 5 h of insulin deprivation, liver D2- and D3-deiodinase messengers decreased or increased, respec-tively. Thus, insulin is another partner involved in bird liver T4 metabolism (Darras et al., 2006 in Dupont et al., 2008). The IGF system also appears partly insulin dependent. Liver IGFBP1 messenger (microarray and qRTPCR measure-ments) and plasma IGFBP1 protein levels increased after insulin immuno-neutralization (plasma IGF1 was not mea-sured). Microarrays also revealed that liver IGFBP2 and muscle IGF1R messengers increased or decreased, respec-tively (similar impairments are found in diabetic patients; Paalsgard et al., 2009 in Simon et al., 2012). In another study (Nagao et al., 2001), 2 day fasting decreased plasma IGF1 and increased IGFBP2 messenger in the liver and giz-zard of white leghorn chickens (but not in brain or kidney); exogenous insulin further decreased plasma IGF1 levels and largely inhibited liver and gizzard IGFBP2 messen-ger (6 h re-feeding also inhibited liver and gizzard IGFBP2 messenger but accordingly increased plasma IGF1 levels).

27.6.4 Insulin and Glucose Metabolism

The severe and rapid hyperglycemia induced by insulin deprivation in fed chickens certainly depends upon multiple changes in glucose metabolism in several tissues. In liver, where glucose transport is not insulin dependent, glucose utilization was probably greatly depressed as suggested by the drop in liver GK messenger and protein contents (see the earlier discussion about the existence and the regulation of liver GK activity). In mammals, other insulin-dependent mechanisms are inhibition of liver gluconeogenesis and glucose production. Gluconeogenesis is considered to escape insulin control in chicken liver (Section 27.1). In our transcriptome study, 27 messengers from the glycolysis–gluconeogenesis or pyruvate metabolism pathways were differentially expressed in liver following insulin depriva-tion, including G6PC2 (glucose 6-phosphatase catalytic, 2) messenger, which increased, suggesting that liver glucose production may also be insulin inhibitable in chicken liver. Unfortunately, this was not assessed in euglycemic–hyper-insulinemic clamp experiments performed on chickens (Chou and Scanes, 1988; Hamano, 2006). Indirect evidence (prolonged fasting and corticosterone experiments) sug-gests that insulin also inhibits kidney gluconeogenesis by inhibiting cytosolic phosphoenolpyruvate carboxykinase activity (Bisbis et al., 1994a,b and quoted references).

Insulin greatly stimulates glucose uptake in muscles and adipose tissues of mammals by recruiting insulin-sensitive

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Glut-4 transporters at the cell surface from an intracellu-lar pool (Figure 27.3). So far, no Glut-4 glucose transporter has been identified in chickens or house sparrows (Passer domesticus) as protein or messenger. This may be the con-sequence of the lack of cross-reactivity of antibodies or the absence of Glut4 in chicken or birds since no Glut-4 homolog sequence is found in the chicken Ensembl data-base (an ortholog gene to the human Glut-4 (SLC2A4) gene is, however, described in the anole lizard and several fish species). Functional and immunoreactive Glut-4 trans-porters have been observed in duck leg muscle (Thomas-Delloye et al., 1999 in Dupont et al., 2009). Only chicken Glut-1, -2, -3, and -8 have been characterized as messen-gers and/or proteins (Carver et al., 2001; Seki et al., 2003; Kono et al., 2005; Swaezea and Braun, 2006 in Zhao et al., 2012; Humphrey et al., 2004). It has been hypothesized that Glut-8 (encoded by the SLC2A8 gene) may act in chickens as Glut-4 does in mammals (Kono et al., 2005; Zhao et al., 2012). In vivo, exogenous insulin increases glucose uptake moderately in some muscles but not in adipose tissue (Tokushima et al., 2005). (Surprisingly, insulin stimulated or inhibited 3H-2DG uptake in liver and brain, respectively.) Thus, an insulin-sensitive glucose transport is present in chicken; however, it requires extensive characterization.

Currently, an increasing number of studies report quan-titative trait loci (QTLs) as controlling growth and/or body composition and some physiological parameters in various avian species. In several cases, these QTL regions harbor glucose- and/or insulin-dependent genes. This aspect can-not be reviewed here, but some references can be found in Nadaf et al. (2009).

27.6.5 Insulin–Glucagon and Lipid Metabolism

Several transcription factors controlling mRNA expres-sion of lipogenic enzymes also appear insulin-sensitive in chicken liver in vivo (SREBP1, PPAR-gamma and THRSP alpha (spot-14), and FASN messengers; Dupont et al., 2008). Carbohydrate-responsive element-binding protein (ChREBP, encoded by the MLXIPL gene) is another tran-scriptional factor involved in glucose metabolism, gluco-toxicity, and/or lipogenesis in liver, adipose tissue, and pancreas in mammals (Leclerc et al., 2012). ChREBP is also activated in parallel with lipogenesis in chicken liver (Proszkowiec-Weglarz et al., 2009). In our insulin neu-tralization experiment, several other enzyme messengers involved in lipogenesis or fatty acid oxidation decreased or increased, respectively, following insulin deprivation (Simon et al., 2012), extending the role of insulin in chicken liver lipid metabolism. In abdominal fat tissue, 5 h of fast-ing altered the level of 2016 messengers. Plasma glucagon increase certainly accounts for a major part of these changes (Ji et al., 2012). Insulin neutralization in fed chickens altered

PART | V Endocrine Theme

the expression of only 92 genes, 72 of which were also dif-ferentially expressed by 5 h of fasting. All genes that were affected by both treatments changed in the same direction (up- or downregulated in both groups). Most messengers that were specifically altered by insulin neutralization (but not by fasting) were either poorly or not described; however, glucagon precursor (class B-preproglucagon) and gluca-gon receptor precursor were in this category and were up- and downregulated, respectively. A local insulin-dependent paracrine glucagon regulation, which needs full character-ization, is suggested in chicken adipose tissue. In general, a contribution of insulin to adipose tissue development and metabolism (in addition to its control of liver lipogenesis) can be expected, despite the very atypical feature exhibited by chicken adipocytes.

Differentiation of chicken adipocyte precursors (ob-tained from the stromal vascular fraction of fat tissue) or transdifferentiation of fibroblasts, epithelial oviduct, or pri-mordial germ cells into adipocytes has been successfully performed using media rather similar to those used for mam-malian adipocyte differentiation (Khuong and Jeong, 2011; Li et al., 2010; Liu et al., 2010; Ramsay and Rosebrough, 2003). Insulin is required but unable to support differentia-tion alone; IGF1 or T3 are unable to replace insulin (Ramsay and Rosebrough, 2003). Chicken serum and dexamethasone can be deleted, provided oleic acid is introduced into the differentiation medium. Unless oleic acid exerts a spe-cific effect, this supports the conclusion that to overcome their intrinsically low lipogenic activity and achieve a full adipocyte phenotype, chicken adipocyte precursors need exogenous fatty acids (Liu et al., 2009; Matsubara et al., 2005, 2008). Several transcription factors are involved in adipogenesis: C/EBPalpha, PPARgamma, and SREBP1 in chicken (Liu et al., 2009, 2010; Sato et al., 2009; Wang et al., 2008b) and duck (Xiong et al., 2010).

In adipocytes of mammals, lipolysis is under the control of several lipases, including the adipose triglyceride lipase, encoded by the PNPLA2 gene, and the hormone-sensitive lipase (HSL), encoded by the LIPE gene (Zechner et al., 2009). HSL (LIPE) disruption in normal or ob/ob mice leads to several changes in the endocrine pancreas and adi-pocytes that resemble the chicken feature (see Osuga et al., 2000; Wang et al., 2001; Haemmerle et al., 2002 quoted in Zechner et al., 2009; Mulder et al., 2003; Harada et al., 2003; Sekia et al., 2009). Although not identified in chicken, the LIPE gene is present in the anole lizard and several fish. If the LIPE gene is really missing in chicken, some of the chicken physiological peculiarities may be due to this.

27.6.6 Insulin and Protein Metabolism

In addition to its general effects on protein metabolism (see Section 27.1; Tesseraud et al., 2007a), insulin acts in the ubiquitin–proteasome proteolytic pathway. It inhibits the

Chapter | 27 Endocrine Pancreas

expression of atrogin-1 messenger in quail muscle (QT6) fibroblasts (Tesseraud et al., 2007b) and in vivo in chicken muscle (Dupont et al., 2008). Following insulin immu-noneutralization, several messengers coding for enzymes of the oxidative pathway decreased. This pathway is also altered in muscle of human diabetics (in Simon et al., 2012).

27.6.7 Insulin and Gene Expression

In mammals, insulin stimulates the translation of preex-isting mRNA and coordinates (stimulates or inhibits) the expression of multiple genes in various tissues. Multiple insulin responsive elements (IRE) have been identified on target genes (other IREs are likely to be identified; Mounier and Posner, 2006). To our knowledge, the characteriza-tion and the functionality of IREs, as such, are still miss-ing in chickens. The fact that several transcription factors or enzymes involved in lipid metabolism are also insulin sensitive in chickens (as discussed in this chapter) suggests that major IREs have been conserved or have co-evolved in chickens. A total of 1573 and 1225 signals were altered by insulin deprivation in liver and muscle, respectively, includ-ing several transcription factors. Several messengers have been associated with energy expenditure, insulin resistance, metabolism sensing, obesity, and/or diabetes in mammals (see Table 3 in Simon et al., 2012). The chicken elongation factor-2 (EF-2) gene, a major factor in protein synthesis in mammals, is also under the control of insulin and 8-bromo-cAMP or phorbol ester (Lim and Kim, 2007). Insulin depri-vation in fed (but not fasting) chickens decreased EGR1 (early growth response 1) messenger in liver and muscle but not in adipose tissue. However, TOE1 (target of egr 1, member 1) was also specifically decreased by insulin depri-vation and not by fasting in adipose tissue (Ji et al., 2012). EGR1 target genes are largely unknown, particularly in the field of metabolism.

Finally, microRNAs (miRNAs) represent another mechanism in the regulation of gene translation. At least miR-196 and miR-200b showed significant and opposite changes in response to insulin neutralization (an increase in liver versus a decrease in muscle; Bouhallier, Rouault, and Simon, unpublished data). Gene targets of miR-196 and miR-200b need to be identified; so far, miR-196 has been involved in chicken embryo development (McGlinn et al., 2009) and miR-200b in chicken liver lipid metabolism (Hu et al., 2012).

27.7 EXPERIMENTAL OR GENETICAL MODELS

Table 27.1 summarizes results about fasting plasma glucose–insulin relationships, glucose tolerance, insulin sensitivity, and fattening in various experimental or genetic chicken models. Most of these results have been discussed

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in previous publications (the effect of prolonged fasting has also been investigated, as discussed in this chapter). The table is shown to support some general considerations. It should be noted that in all these experiments, the feeding pattern of chickens has been consistently synchronized with light–dark cycles. In these conditions, as in mam-mals, the fasting plasma glucose level appears as an indica-tor or a central node, which reflects or integrates general metabolism. Although the primary mechanism is unknown, changes in glucose–insulin balance and/or insulin sensi-tivity are associated with changes in fattening. In contrast to obese mammals, fat chickens have low plasma glucose and do not develop insulin resistance. Conversely, chickens from a low-glucose line are fatter than their counterparts. Only the corticosterone model behaves like obese mammals (showing higher plasma glucose and impaired insulin sen-sitivity). As discussed here, changes in insulin signaling are present in the models thus far investigated. Changes in glu-cose tolerance are not consistently associated with changes in body composition: in a commercial broiler cross, chick-ens exhibiting fast or slow plasma glucose regulation at 2 and 4 weeks of age following an oral glucose load did exhibit fast or slow glucose disposal at 8 weeks of age but did not differ in fasting or glucose-induced plasma insulin levels and body composition, which remains unexplained (Simon, 1980). One can hypothesize that, as in FL chick-ens, enhanced glucose-induced insulin levels are required to increase fattening without necessarily leading to insulin resistance. In vitro, perfused pancreas from fat chickens released less insulin during the first phase in response to glucose, or to arginine or acetylcholine in the presence of glucose, than pancreas from lean chickens, which remains unexplained in Rideau (1988). In general, chickens appear to be controlled within a much narrower range for their body composition than mammals. Whether the endocrine pancreas of birds plays a part in this remains an open ques-tion and a challenge for the future.

27.8 SUMMARY AND CONCLUSION

Previous reviews, summarized in the Section 27.1, revealed several features of the chicken endocrine pancreatic physiol-ogy. The present chapter is focused on glucagon and insulin; it highlights new knowledge about chicken embryo devel-opment, glucagon (and glucagon-related peptides), bird insulins, glucagon and insulin release, glucagon and insu-lin receptors, and major general effects of these hormones. Many results highlighted in this review point out further chicken peculiarities at the level of endocrine pancreatic physiology. Chickens (and birds in general) developed sev-eral species specificities during evolution. The most original ones concern the refractoriness of glucose-induced insu-lin release and the insulin signaling in muscle and adipose tissue. One unidentified inefficient metabolic step makes

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glucose a poor insulinotropic nutrient in chickens. Although several elements of insulin signaling pathways have not been investigated yet, several steps of the IR cascade appear insensitive in muscle; the feature is even worse in adipose tissue. In contrast, IR signaling appears “normal” in chicken liver. Revealing mechanisms or steps accounting for these chicken peculiarities may provide important information, including for human diabetes pathology. Glucagon physiol-ogy appears more complex in chicken than in mammals: the glucagon gene encodes several transcripts, it is expressed in adipose tissue, two types of glucagon receptors are present, and an insulin-dependent paracrine glucagon regulation is suggested in adipose tissue. In the fed status, insulin is coor-dinating the endocrine balance to permit the development or the maintenance of an active anabolism. From a practical point of view, finding the way to favor pleiotropic anabolic actions of insulin in muscle and minimize liver lipogenic and adipogenic actions of insulin would be highly profitable for human nutrition and poultry production. This chapter also pointed out many shortcomings or new areas to explore. In short, the secretory “machinery” of A- and B-islets needs to be fully characterized. Chicken is the unique model to pro-vide such possibilities. In addition, nothing is known about the physiological actions of GLP-2, oxyntomodulin, mini-glucagon, glicentin, or GRPP (gastrin-releasing pancreatic peptide) in chicken. Full characterization of IR signaling and the subsequent steps up to the control of gene expres-sion certainly also offer large areas for future research. Finally, chicken glucose transporters (and their presence and functionality) need to be characterized. Although chicken is inconvenient because it requires the development of tech-nical tools for each area to explore, it remains an almost unexplored experimental animal model of large interest for comparative endocrinology and physiology.

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