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Leptin is an anorexigenic hormone of 16 KDa, mainly produced by the adipose tissue (Zhang et al., 1994). Leptin plays an important role in the central regulation of energy balance and is released by adipose tissue in proportion to the amount of lipid stores (Maffei et al., 1995; Ostlund et al., 1996). It acts on hypothalamic receptors (Campfield et al., 1995), decreasing food intake and increasing energy expenditure (Himms-Hagen, 1999; Ahima and Flier, 2000). Leptin participates also in multiple cellular and physiological processes mainly linked to the control of body weight such as gastric emptying, thermogenesis, nutrient absorption in the gut, lipolysis, and metabolic fuel oxidation (Reidy and Weber, 2000; Muoio and Dohm, 2002). The leptin receptor (OB-R) is a member of the cytokine Class I receptor superfamily and was cloned from brain (Tartaglia et al., 1995). Based on the length of the intracellular domain, the OB-R has one long isoform (OB-Rb) and four short isoforms (OB-Ra, c, d, f), (Cinti et al., 2001). Another soluble isoform (OB-Re) lacks the transmembrane domain and may be involved in leptin transport in the blood. Apart from the original tissues from which leptin and its receptor were cloned, expression of these proteins has been found also in many other tissues, including gastrointestinal tract (Bado et al., 1998; Cinti et al., 2001; Cammisotto et al., 2005), placenta (Masuzaki et al., 1997), skeletal muscle (Wang et al., 1998), and mammary epithelium (Casabiell et al., 1997), thus suggesting a wider role of this protein (Sanchez et al., 2005). Moreover, the expression of leptin was detected in lung of gray and harbor seal suggesting its potential role in marine mammal respiratory physiology (Hammond et al., 2005). Interestingly, leptin mRNA and protein have been found in the fundic epithelium of the rat (Bado et al., 1998), and leptin and its receptor are expressed in human gastric epithelial cells (Sobhani et al., 2000). Gastric leptin is secreted both in the plasma as hormone-receptor complex and in the gastric juices that reach the duodenum notwithstanding the acid environment (Buyse et al., 2004; Cammisotto et al., 2005, 2006). Recent reports have described leptin immunoreactivity in intestinal enterocytes of duodenum, jejunum, ileum, and colon of the mammals (Cammisotto et al., 2005; El Homsi et al., 2007), in the lining epithelium and in selected visceral fibers of the gastrointestinal tract of nonmammalian vertebrates (Muruzàbal et al., 2002; Neglia et al., 2008; Gambardella et al., 2010; Russo et al., 2011). Moreover, in mammals, OB-R was detected in some elements of the enteric nervous system (Liu et al., 1999) and here, the bond with its ligand causes numerous functions. In fact, leptin is involved in intestinal lipid handling (Morton et al., 1998) and sugar adsorption (Lostao et al., 1998; Pearson et al., 2001), and, at least in cats, controls intestinal motility throughout a long-loop reflex involving intestinal vagal afferent fibers (Gaigè et al., 2003).
The morphology of the gastrointestinal tract of vertebrates evolved following specific metabolic demands and individual requirements for processing, distributing, and absorbing nutrients, and eventually regulates body weight (Williams et al., 2001; Szekely et al., 2010). In a series of previous studies, our research group investigated the relationship between the expression of leptin and the different anatomical organization of the nonmammalian vertebrates, including two teleostean species (bass and goldfish) with different adaptive morphological organization of the gastroenteric tract and feeding modalities (stomach-containing and stomach-less teleosts) (Russo et al., 2011). The presence and distribution of leptin has been studied also in the water buffalo (Russo et al., 2009) and in the pig (Russo et al., unpublished). Comparison of the results suggested that leptin immunoreactivity can be found in the parts of the gastrointestinal tract that carry out an equivalent digestive function (e.g., in the stomach of bass and in anterior intestine of goldfish; in the abomasum of water buffalo and stomach of the pig). Thus the expression of peripheral peptides in these vertebrates is apparently related to the function and not to the gross morphology of the organ. Consequently, the topographical localization of leptin immunoreactivity shows a close relationship with specific organization of the gut. No data are present in the literature relative to the leptin expression in the gut of monogastric and polygastric marine mammals, like sea lions and dolphins that share the same environment but have evolved different strategies for food intake and processing.
With this study, we intend to provide further insights on the neuroendocrine regulation of the digestive postdiaphragmatic functions by using an immunological approach to, the expression of leptin, in the gastrointestinal tract of two species of marine mammals, a poligastric cetacean (Tursiops truncatus) and a monogastric pinniped (Otaria flavescens).
MATERIALS AND METHODS
Animals and Tissue Preparations
For the present study we used a series of samples of the gastrointestinal tract of the South American sea lion Otaria flavescens (two specimens) and the bottlenose dolphin Tursiops truncatus (three specimens) stored at the Mediterranean marine mammal tissue bank of the University of Padova (http://www.mammiferimarini.sperivet.unipd.it/eng/index.htm). The samples used are listed in the Table 1.
Table 1. Specimens and samples used for histochemical dyes and immunohistrochemistry
Otaria flavescens ID 128 (♂)
Otaria flavescens ID 169 (♂)
Tursiops truncates ID 107 (♂)
Tursiops truncatus ID 110 (♂)
Tursiops truncates ID 139 (♂)
First chamber or forestomach
Second chamber or mainstomach
Third chamber or pyloric stomach
The samples were removed within a few hours after death of the animals. Some fragments were frozen and stored at −80°C, while others were fixed in 10% buffered formalin and later embedded in paraffin. Sections were serially cut into 8-μm-thick transversal sections, placed on slide glasses. For anatomical studies one every ten histological samples were stained by common histochemical dyes (hematoxylin-eosin and alcian-PAS stainings) for morphological orientation, while the other sections were used for immunohistochemistry.
The expression and distribution of cromogranin A (Chr A), a marker of neuroendocrine cells (Deftos, 1991), polypeptide gene product (PGP 5), a marker of enteric nervous system (Krammer et al., 1993), and leptin in the gastroenteric tract of Otaria flavescens and Tursiops truncatus were studied by immunohistochemistry. The sections were dewaxed and incubated with 0.3% hydrogen peroxide for 30 min at room temperature (RT), to block endogenous peroxidase activity. The sections were then rinsed in 0.01 M phosphate-buffered saline (PBS), pH 7.4, for 15 min and subsequently incubated for 20 min at RT with normal goat serum. Normal serum and the other components of the immunological reaction were contained in the Vectastain Elite ABC kit (PK 6101; Vector Laboratories, CA). In the specific step, polyclonal antibodies raised against ChrA (20086, Immunostar, WI), PGP 9.5 (E3344, Spring Bioscience, CA), and leptin (A-20, sc-842 Santa Cruz Biotechnologies, CA) were diluted 1:1,000; 1:500; and 1:300; respectively. After incubation with primary antisera the sections were rinsed in PBS for 15 min and incubated for 30 min at RT with biotinylated goat anti rabbit IgG. Subsequently, the sections were rinsed in PBS for 15 min and then incubated for 30 min at RT with avidin-peroxidase complex. Peroxidase activity was detected using a solution of 3-3′ diaminobenzidine tetrahydrocloride (Sigma, St. Louis, MO) of 10 mg in 15 mL 0.5 M Tris buffer, pH 7.6, containing 0.03% hydrogen peroxide.
Double immunohistochemical staining using two primary antisera raised in the same species, that is, rabbit, was performed according to the method described by Wessel and McClay (1986) which consisted of the use of fluorochrome-conjugated Fab fragments to avoid crosstalk of subsequent antisera. In this case, dewaxed, rehydrated, blocked sections were rinsed in 0.01 M PBS (pH 7.4) containing 0.2% Triton X-100 and 0.1% bovine serum albumin. The sections were then incubated with PGP 9.5 antiserum (1:50 in 1:5 normal goat serum for 48 h at RT). The sections were then washed in PBS and incubated with GAR-Fab fragment conjugated to FITC fluorochrome (1:30 for 2 h at RT; 713-095-147; Jackson lab., West Grove, PA). Thereafter, the sections were rinsed in PBS and incubated with rabbit anti leptin (1:20 for 48 h at RT). After having been rinsed in PBS, the sections were treated with affinity-pure donkey anti-rabbit IgG conjugated to TRITC fluorochrome (1:50 for 2 h at RT; 711-025-152; Jackson). Finally, the sections were washed with PBS, mounted with glycerin diluted with PBS 1:1.
The immunostained sections were photographed using a Leica DMRA2 microscope (Leica, Wetzlar, Germany) equipped with a DC300F digital camera.
Controls were obtained by absorbing each primary antiserum with an excess of the relative peptide (100 mg of peptide per mL of diluted antiserum; Ob hBA-147/sc-4912; Santa Cruz) and by substituting primary antisera with PBS or normal serum in the specific step. Archival samples of male Wistar rat stomach (Harlan, Italy), known to be positive to Chr A, PGP 9.5 and leptin, were used as positive reference controls.
Western Blot Analysis
Proteins from South American sea lion and bottlenose dolphin gastrointestinal tract and mouse stomach were extracted with Lysis buffer (50 mM NaCl, 25 mM Tris [pH 8.0], 0.5% NP40, 0.1% SDS, 1 mM protease inhibitors [Roche], and 1 mM phenyl-methylsulfonyl fluoride). Samples were spun at 13,000 rpm for 20 min, and supernatants were collected. Protein concentrations were determined with the Bio-Rad dye protein assay. Samples were boiled at 98°C for 10 min, frizzed-and-thawed, and boiled again at 98°C for 10 min. The proteins were separated on a 15% SDS–polyacrylamide gel electrophoresis with 4% stacking gel in 1% Tris–glycine buffer (0.025 M Tris, 0.192 M glycine, and 0.1% SDS [pH 8.3]) in a miniprotean cell (Bio-Rad) at 130 V for 2 hr. The separated proteins were electrotransferred onto a polyvinylidene fluoride membrane with transfer buffer (25 mM Tris base, 0.2 M glycine, and 20% methanol [pH 8.5]) in a minitransfer cell (Bio-Rad) at 100 V at 4°C for 1 hr. Membranes were incubated at 4°C for 1 hr in blocking buffer containing 1% PBS and 0.05% Tween 20 with 5% dried nonfat milk and then were probed with an polyclonal antibody that recognizes a synthetic fragment of human leptin overnight at 4°C. This was followed by incubation with a goat secondary anti-rabbit IgG (1:5,000) for 1 hr at RT. Signals were detected by chemoluminescence with the Pico Enhanced Chemiluminescence Kit (Pierce Chemical). A prestained molecular-weight ladder (Fermentas) was used to determine protein size.
To ensure specificity, preabsorption of leptin antibody with its relative control peptide was performed before western blotting.
The gastrointestinal tract of Otaria flavescens is typical of monogastric carnivores. The stomach has a single chamber and intestine is divided into small and large intestine. The duodenum is characterized by a typical C-shape and the end of small intestine and the start of colon is marked by the presence of cecum diverticulum. The histological features of the all gastrointestinal segments are similar to carnivores ones.
The gastric complex of the bottlenose dolphin is made up by three gastric compartments: forestomach or first chamber (Fig. 1F), main stomach or second chamber (Fig. 1M) and pyloric stomach or third chamber (Fig. 1P), linked to the previous stomach by two connecting chambers or channels (Fig. 1 1,2). The forestomach is lined with stratified squamous epithelium which is continuous with the esophageal epithelium. The musculature of the forestomach is prominent and thick, and displays neither glands nor muciparous cells. The communication between the forestomach and main stomach is relatively wide and open. The main stomach is the homologous of gastric fundus of monogastric mammals. Lined with columnar epithelium, its mucosa is rich in tubular gastric glands and contains muciparous, parietal, and chief cells. The muscular lining of the main stomach is relatively thin. The connecting chambers are a narrow, twisted sphincteric passage, provided with a valve. Their epithelial lining is thin and contains pyloric glands. The third chamber or pyloric stomach presents a simple structure lined with a relatively thin columnar epithelium. It contains muciparous cells organized into pits or pyloric glands. The muscular wall of the pyloric stomach is thinner than any of the other compartments. The third stomach communicates with the duodenal ampulla (Fig. 1DA) by means of the heavily muscular pyloric sphincter.
The intestine showed no distinctive macroscopical characteristics and histological examination revealed a constant and repetitive morphology in the proximal, middle, and distal tracts. The mucosal layer of all intestinal tracts rises in folds and contains small crypts of Lieberkhun. These are not ramified and present numerous goblet cells. The intestine is lined with columnar epithelium with a basal nucleus and its mucosa contains glands. The muscular wall is thin. In the distal tract there are not intestinal villi.
Because primary structures of cetacean and pinniped peptides are unknown and crossreactivity between different peptides containing the same antigenic sequences cannot be excluded, the immunoreactive material is properly referred to as “peptide-like” immunoreactivity.
South American sea lion.
Numerous Chr A-like-immunoreactive (ir) cells were localized in the mucosal epithelium of the stomach (Fig. 2A), duodenum (Fig. 2B), and jejunum. In detail, these cells were found at the basis of the mucosal folds in the stomach (Fig. 2A) and in the glands of the other immunopositive gastrointestinal segments (Fig. 2B). Chr A-like-ir cells showed a focal distribution being grouped in small clusters of 5–10 elements each. Their shape was round (Fig. 2A) or irregular (Fig. 2B). PGP 9.5-like-ir neurons and nervous fibers were detected in the submucosal (Fig. 2C,D) and myenteric plexuses of the all gastrointestinal segments investigated in Otaria flavescens. The nervous fibers were thin, often varicose and travelled independently or grouped in small bundles (Fig. 2C). PGP 9.5-like-immunopositive neural bodies were abundant and either isolated or clustered (Fig. 2D). The -ir materials filled the entire cytoplasm and showed different intensity of staining. Some leptin-like-ir cells were found in the pit of gastric mucosal folds (Fig. 3B) and very few were present also in the mucosal glands of duodenum. These cells were round (Fig. 3B) or elongated in shape, showed the typical morphology of endocrine cells. Several leptin-like-ir nurons were identified in the submucosal (Fig. 3C) and myenteric plexuses of the stomach.
By double immunostaining, some PGP 9.5- and leptin-like ir neuronal structures were detected both in submucosal than in myenteric plexuses of the stomach.
Several Chr A-like-ir cells were identified in the main stomach; pyloric stomach (Fig. 4 A,A1); proximal and middle intestine (Fig. 4B) but not in the forestomach. PGP 9.5-like-ir neurons and nervous fibers were detected in the submucosal and myenteric plexuses (Fig. 4C) of the all gastrointestinal segments investigated.
Many leptin-like-ir cells were identified in mucosal layer of the second chamber. They were located in the lower half of the glands (Fig. 5A,B). No positive cells were found in the mucosal epithelium of the other gastric compartments and intestinal tracts.
Many leptin-like-ir neural bodies and fibers were scattered in the submucosal and myenteric plexuses of the main stomach (Fig. 5C), pyloric stomach (Fig. 6A), proximal, and middle intestine (Fig. 6B,C1). Immunopositive neurons were grouped (Figs. 5C, 6B,C1) or isolated (Fig. 6A) and always showed a strong intensity of the staining. Nerve fibers traveled alone or in small bundles and were sometimes seen in perivascular localizations (Fig. 6B), namely in small arteries and arterioles.
Double immunostaining showed leptin-like immunoreactivity coexisted with PGP 9.5-like one in numerous neurons and fibers located both in submucosal than in myenteric plexuses of main stomach (Fig. 7A,B), pyloric stomach, proximal, (Fig. 7C,D) and middle intestine.
Western Blot Analysis
Western blot analysis was carried out on lysates of all gastrointestinal tract of bottlenose dolphin (forestomach, main stomach, pyloric stomach, proximal, and middle intestine) and South American sea lion. The results of the western blot analysis showed the canonical mammals 16 kDa band of the protein for all gastrointestinal tracts with the exception of bottlenose dolphin forestomach (Fig. 8). The specificity of the response was confirmed by preincubation of the leptin antibody with its respective blocking peptide. There was no expression of leptin in these preparations, whereas the presence of the proteins was detected in a mouse stomach homogenate that was used as positive control (data not shown).
Our study confirms previous observations (Bryden, 1972; Green, 1972; Mead, 2008) on the anatomy of the gastrointestinal tract in seals and dolphins. The stomach of the South American sea lion is similar to that of other carnivore, while the gastric complex of the bottlenose dolphin is divided into different chambers (Harrison et al., 1970; Gaskin, 1978; Mead, 2007, 2008), and composed of a forestomach or first chamber, a main stomach or second chamber and a pyloric stomach or third chamber connected by minor chambers or channels. Dolphins cannot chew their food, and the complexity of their stomach with a large and distinct forestomach possibly reflects the necessity to grind the ingested fish or squid. The large passage between the first and the second chambers suggests frequent exchanges of materials between the two stomachs before progression into the pyloric chamber, protected by a twisted connection with sphincteric walls and a valve.
It is also worth noting that the morphology of the intestine of the bottlenose dolphin is very uniform during its entire length, and the gut cannot be divided into separate parts based on macroscopic characteristics or structure. The duodenum is conventionally indicated as a separate portion due to the presence of the hepatopancreatic duct. The definition given here of proximal, middle and distal intestine is indicative of the position of the sampled tract relative to the whole length of the gut, but we were not able to detect any structural variation.
The patterns of distribution of Chr A-like protein, a marker of neuroendocrine cells (Deftos, 1991), PGP 9.5-like protein, a marker of enteric nervous system (Krammer et al., 1993) and leptin-like protein, are described here for the first time in the digestive apparatus of marine mammals. In previous studies, the presence of argentaffin cells were found in the first chamber of poligastric sea mammals (Harrison, 1970; Gaskin, 1978) and in these cells serotonin was coexpressed (Gaskin, 1978). Also, in the bowhead whale (Balaena mysticetus) enteroendocrine cells were identified within the mucous gland of cardiac region, connecting channels, pyloric chamber, and cranial duodenum (Tarpley et al., 1987). In our experimental series, Chr-A-like immunopositive endocrine cells were scattered in the mucosal layer of stomach and intestine of Otaria flavescens as well as in the epithelial mucosa of main stomach, pyloric stomach, and intestine of Tursiops truncatus. Furthermore, PGP 9.5-like ir neuronal bodies and fibers were identified in the submucosal and myenteric plexuses of the all gastrointestinal segments of both marine mammals. Leptin-like immunopositive cells were found also at the basis of mucosal folds of the stomach and in the mucosal layer of the proximal intestine of the South American sea lion and in the glands of main stomach of the bottlenose dolphin. Moreover, numerous leptin-like ir neurons and fibers of submucosal and myenteric plexuses were detected in the gastrointestinal tracts of both species. These elements of the enteric nervous system were also positive to PGP 9.5-like protein.
The gastric mucosa appears to be the only tissue secreting leptin in an exocrine rather than endocrine mode in mammals (Cammisotto et al., 2005, 2006). In fact both in rats and humans, leptin-positive cells were localized in the lower half of the gastric mucosa (Mix et al., 2000; Cammisotto et al., 2005). Secreting cells belong to two different populations: the most frequent type is similar to pepsinogen-secreting chief cells; the second cell type displays the morphology characteristic of endocrine cells (Cinti et al., 2000; Cammisotto et al., 2005, 2006). Our results suggest that leptin-like ir gastric cells can be classified into two types also in marine mammals, but these two cell population do not coexist in the same species. Leptin-like ir cells in the stomach of Otaria flavescens were few, showed the typical morphology of endocrine cells, and were similar to Chr-A-like ir elements; on the other hand, leptin-like ir cells found in the mainstomach of Tursiops truncatus closely resembled exocrine type cells. The presence of leptin-like-ir cells in the main stomach of the bottlenose dolphin suggests a rapid leptin secretion by chief cells after the onset of food intake in accord to the short-term control of food intake (Cammisotto et al., 2005). Thus an involvement of leptin-like peptides in the stimulation of mucus secretion and in anti-acid gastrointestinal protection could be possible also in this cetacean species, (Plaisancie et al., 2006). Vagal stimulation or secretion of hormonal factors (CCK in rats and secretin in humans) induce leptin release into gastric juices linked to a protein of high molecular weight (Bado et al., 1998; Attoub et al., 1999; Cinti et al., 2000; Sobhani et al., 2000; 2002, Guilmeau et al., 2003). Some of the stomach-derived leptin is not fully degraded by proteolysis, and therefore reaches the intestine in an active form and may act in the control of specific intestinal functions. Recent reports have described leptin immunoreactivity in enterocytes of duodenum, jejunum, ileum, and colon of mammals (Cammisotto et al., 2005; El Homsi et al., 2007). We have detected very few leptin-like-ir cells only in the proximal intestine of the South American sea lion. The majority of these latter elements occupies the outer epithelial layers and faces the lumen. Their location indicates a possible paracrine/autocrine or even endocrine action of leptin directly on intestinal tissues in sea lions, as in land mammals (Guilmeau et al., 2003; Cammisotto et al., 2005, 2006).
The presence of leptin-like immunopositive neurons and fibers in the enteric nervous system of both species of marine mammals is in agreement with what is presently known in non mammalian vertebrates (Muruzàbal et al., 2002; Bosi et al., 2004; Neglia et al., 2008; Russo et al., 2011). There are no functional data relative to a specific role of leptin in the digestive physiology of marine mammals. However a series of studies performed in mammals (Morton et al., 1998; Lostao et al., 1998; Pearson et al., 2001; Guilmeau et al., 2003; Yarandi et al. 2011), showed that leptin-like peptides are active in the regulation of motility and the absorption of nutrients. Leptin has been recently reported to excite the submucous and myenteric neurons of the guinea pig (Reichardt et al., 2011) and to control cat intestinal motility by a long-loop reflex involving intestinal vagal afferent fibers (Gaigè et al., 2003). The role of leptin in the gut of marine mammals, and especially in the complex gastric chambers of dolphins, cannot be ascertained on the basis of the present study alone. However, the diffuse presence of the protein in the gut may suggest a function in the regulation of food progress in the digestive system.
The morphology of gastrointestinal tract reflects the metabolic demands of the animals and, at least in other vertebrates, is related to the modification of expression of peripheral leptin-like peptides. Here we note that although the monogastric South American sea lion and the poligastric bottlenose dolphin have a very different anatomical organization of the gut, the distribution of leptin-like peptides is similar and affects compartments that are active in digestive physiology (stomach of the seal lion; main and pyloric stomachs of the dolphin; intestine of both) but not those parts (forestomach) of the dolphin gut that play only a mechanical role in food processing. We further note here that both species have a high digestive rate, and this latter factor may be linked to a diffuse distribution of leptin-like -ir cells. Further investigations involving blood sampling and controlled dietary supplement may help to study the role of leptin-like peptides in the regulation of food intake, body weight and energy balance in marine mammals.