Leptin Orally Supplied to Neonate Rats Is Directly Uptaken by the Immature Stomach and May Regulate Short-Term Feeding
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内分泌学杂志 2005年第6期
Laboratorio de Biología Molecular, Nutrición y Biotecnología, Departamento de Biología Fundamental y Ciencias de la Salud, Universitat de les Illes Balears, E-07122 Palma de Mallorca, Spain
Address all correspondence and requests for reprints to: C. Picó, Department of Biología Fundamental y Ciencias de la Salud, Universitat de les Illes Balears, Cra. Valldemossa Km 7.5, E-07122 Palma de Mallorca, Spain. E-mail: cati.pico@uib.es.
Abstract
Although leptin is a hormone mainly produced by the adipose tissue, it is also produced by the gastric mucosa and the mammary epithelium and is present in maternal milk. The effects of milk leptin on the neonate are not known. The purpose of the investigation was to evaluate the short-term effects of the administration of a single oral dose of leptin on 4-d-old rats as well as the effects of chronic supplementation during the lactation period with a daily oral dose of leptin (equivalent to 5 times the amount of leptin ingested normally from maternal milk during the suckling period) on body weight, the gastric leptin system, gastric food content, and thermogenic capacity. Our results show that the administration of a single oral dose of 4 ng of leptin to 4-d-old rats produces a short-term increase in leptin levels in the stomach and serum and a decrease in the weight of the gastric contents. Pups treated with a daily oral dose of leptin during the whole lactation period showed, at the end of the suckling period, compared with controls, lower gastric contents, lower leptin production by the stomach and the sc adipose tissue, and lower thermogenic capacity in brown adipose tissue. We conclude that oral leptin is absorbed by the immature gastric epithelium of the neonate, and this leptin exerts clear biological effects, down-regulating endogenous leptin production and playing a potential role in the short-term control on food intake during the lactation period.
Introduction
LEPTIN IS AN anorexigenic hormone, mainly produced by the adipose tissue (1), which plays an important role in the central regulation of energy balance. This hormone is released into the circulatory system by the adipose tissue in proportion to the amount of lipid stores (2, 3) and acts at the hypothalamic receptors (4), decreasing food intake and increasing energy expenditure (5, 6). Leptin production in nonadipose tissue such as stomach (7, 8, 9), placenta (10, 11), skeletal muscle (12), and mammary epithelium (13) has also been reported, thus extending the role of leptin to various biological functions (5, 14).
Leptin in the human stomach is located in the lower half of the stomach glands, particularly in the pepsinogen granules of chief cells and in a specific endocrine cell-type (8, 9). There is evidence indicating that feeding stimulates the secretion of leptin by the stomach in humans (8, 9) and rats (7, 15), in which a short food intake stimulus (20 min of refeeding) after 14 h of fasting is capable of practically emptying the leptin stores from within the stomach mucosa (15). In addition, gastric leptin shows diurnal variations influenced by food intake rhythms and changes occurring just before the beginning of the feeding period are opposite to those of the orexigenic hormone ghrelin (16). Pepsinogen secretagogues such as cholecystokinin, gastrin, or secretin also induce gastric leptin release in rats (7).
Leptin receptors are present in the human stomach (17, 18, 19) and mouse gastrointestinal tract (20). Leptin can modulate vagal afferent fibers that originate in the gastric and intestinal walls and terminate in the nucleus tractus solitarius (21), providing rapid information to the brain. These data, together with the observed sensitivity of gastric leptin to the nutritional state, have allowed us to suggest a role for gastric leptin in the short-term regulation of food intake, probably acting as a satiety signal (22).
In neonate rats, leptin production by the stomach probably starts at the onset of suckling, although leptin production by the stomach is kept at low levels during the suckling period (23). The change of diet from milk to solid chow is associated with a significant increase in gastric leptin production (23). The presence of leptin in maternal milk and the fact that leptin in nursing rats could be absorbed by the gastric mucosa of neonatal rats (23) and transferred to the infant rat circulation (13) have suggested that leptin supplied from maternal milk could be the main source of leptin in the stomach during the suckling period (23). This exogenous leptin could exert biological effects in neonates at a time in which both the adipose tissue and appetite regulatory systems are immature (21).
Leptin concentrations in human milk vary significantly between people (24, 25), and some authors have found a positive correlation between leptin concentration in milk and maternal plasma leptin levels and adiposity (24). Thus, breast-fed infants nursed by mothers with significant adiposity may be exposed to higher amounts of leptin than infants nursed by lean mothers, and much higher than those fed with infant formulas, which do not have leptin as an ingredient (26). However, the effects of oral intake of leptin on neonate development are not known.
The aim of this study was to evaluate the effects of the administration of an acute oral dose of leptin on food intake in neonatal rats as well as the prolonged effects of the chronic administration of a daily oral dose of leptin, close to physiological levels, during the suckling period on the gastric leptin system and energy balance to determine the importance of leptin supplied by maternal milk during lactation.
Materials and Methods
Animals and experimental design
Study I: study of milk leptin concentration in nursing rats during lactation.
Three-month-old, virgin female Wistar rats were caged with a male rat. Animals were from Charles River Laboratories (Barcelona, Spain). After matching, each female was placed in an individual cage with free access to water and food. Rats were kept in a room with controlled temperature (22 C) and a 12-h light, 12-h dark cycle (light on from 0800 to 2000 h). After delivery, nine dams were used for milk studies. Birth was defined as d 0 of lactation. Milk and serum samples of each dam were collected on d 7, 14, and 21 of lactation. For milk collection nursing rats were separated from their pups for 6 h to guarantee that mammary glands were full of milk. Before milking, dams were exposed to ether, and then milk was obtained from the mammary glands by manual milking and stored frozen at –20 C. A sample of blood was also collected from the end of the tail, stored at room temperature for 1 h and overnight at 4 C, and then centrifuged at 1000 x g for 10 min to collect the serum.
Study II: short-term effect of the administration of a single oral dose of leptin in 4-d-old rats.
Four nanograms of recombinant murine leptin (PeproTech, London, UK) or vehicle (water) were orally administrated with a micropipette to 4-d-old Wistar suckling rats belonging to five different litters. After leptin/vehicle administration, pups were kept with their mothers. Rats were killed by decapitation at different times: time 0, just before leptin administration, and 1 and 4 h after leptin or vehicle administration. A sample of blood was collected and processed as described above (study I) to obtain the serum. The stomach was immediately removed in its entirety, opened, rinsed with saline containing 0.1% diethyl pyrocarbonate (Sigma, Madrid, Spain), frozen in liquid nitrogen, and stored at –70 C until analysis. A lengthway fragment from the stomach was fixed by immersion in 4% paraformaldehyde in 0.1 M phosphate buffer (PB) (pH 7.4) overnight at 4 C for immunohistochemistry studies. Gastric contents were weighed.
Study III: effect of chronic administration of oral leptin during the lactation period in neonatal rats.
At d 1 after delivery, pups from the same litter were randomly distributed into two groups: control group and leptin-treated group (n = 5 in each group). From d 1 to d 20 of lactation and during the first 2 h of the beginning of the light cycle, 20 μl of the vehicle (water) or a solution of recombinant murine leptin (PeproTech) was given orally every day to the pups using a pipette. The amount of leptin given to animals was progressively increased from 1 ng of leptin on d 1 to 43.8 ng of leptin on d 20. This dose of leptin was calculated as 5 times the amount of the daily leptin intake from maternal milk, which was calculated from leptin concentration in milk during the lactation period (obtained from study I) and the estimated total daily milk intake during the lactation period (27). We considered this dose of leptin as close to physiological levels, taking into account the range of variation of milk leptin levels in dams. At weaning, on d 21, pups were killed and the stomach, gastrocnemius muscle, brown adipose tissue (BAT), and different white adipose tissue depots (WAT) (epididymal, retroperitoneal, and inguinal) were rapidly removed. The stomach was opened and rinsed with saline containing 0.1% diethyl pyrocarbonate (Sigma). The gastric contents and BAT and WAT were weighed. All samples were immediately frozen in liquid nitrogen and stored at –70 C until RNA analysis. A lengthway fragment from the stomach was fixed by immersion in 4% paraformaldehyde in 0.1 M PB (pH 7.4) overnight at 4 C. Blood was also collected and processed as described in study I to obtain the serum.
The animal protocol followed in this study was reviewed and approved by the bioethical committee of our university and guidelines for the use and care of laboratory animals of the university were followed.
RT-PCR analysis of leptin mRNA in the stomach and WAT
To determine the expression of leptin in the stomach and WAT, we developed a RT-PCR using the housekeeping gene ?-actin as internal control. Total RNA was extracted using Trizol (Invitrogen, Barcelona, Spain) reagent according to the instructions provided by the supplier; 0.25 μg of total RNA (in a final volume of 5 μl) was denatured at 90 C for 1 min and then reverse transcribed to cDNA using Moloney leukemia virus reverse transcriptase (according to the procedure of Applied Biosystems, Foster City, CA) at 42 C for 1 h, with a final step of 5 min at 99 C in a 2400 thermal cycler (PerkinElmer, Norwalk, CT). Sample was denatured at 94 C for 3 min, and then PCR was carried out using the following parameters: 94 C for 1 min, 58 C for 1 min, and 72 C for 2 min. Thirty cycles were carried out for stomach samples and 23 cycles for the WAT samples. The amplification was finished by an extension step of 10 min at 72 C. Primers for the lep gene were: forward 5'-CCA GGA TGA CAC CAA AAC CCT C-3' and reverse 5'-ATC CAG GCT CTC TGG CTT CTG C-3', and for the ?-actin gene: forward 5'-ACG GGC ATT GTG ATG GAC TC-3' and reverse 5'-GTG GTG GTG AAG CTG TAG CC-3'. The expected size of the products were 316 bp for the lep gene and 164 bp for the ?-actin gene, which were visualized by electrophoresis in a 1.5% agarose gel containing ethidium bromide and verified by using a DNA 100-bp ladder. The bands in the gel were quantified by scanner photodensitometry using the Kodak 1D Image Analysis software (version 3.5 for Windows; Eastman Kodak Co., Rochester, NY). The signal for leptin mRNA was normalized to the signal of the housekeeping gene ?-actin and the results were expressed as the leptin to ?-actin mRNA ratio.
Northern blot analysis of uncoupling protein (UCP)1 and UCP3 mRNA
mRNA levels for UCP1 in BAT and UCP3 in gastrocnemius muscle, BAT, and inguinal WAT were determined by Northern blot. Thirty micrograms of total RNA were denatured with formamide/formaldehyde and then fractionated by agarose gel electrophoresis as previously described (28). The RNA was transferred onto a nylon membrane (Roche, Barcelona, Spain) in 20x saline sodium citrate (SSC) buffer: 1x SSC is 150 mM NaCl, 15 mM sodium citrate (pH 7.0) by capillary blotting for 16 h and fixed with UV light.
The specific mRNA were detected by a chemiluminescence based procedure, using a 30-mer antisense oligonucleotide probe (5'-GTTGGTTTTATTCGTGGTCTCCCAGCATAG-3') for UCP1 detection and a 32-mer antisense oligonucleotide probe (5'-GACTCCTTCTTCCCTGGCGATGGTTCTGTAGG-3') for UCP3, both probes synthesized commercially (Roche), and labeled at both ends with a single digoxigenin (DIG) ligand. Prehybridization was at 42 C for 15 min in DIG-Easy Hyb (Roche). Hybridization was at 42 C overnight in DIG-Easy Hyb containing 34 ng/ml of the oligonucleotide probe. Then hybridized membranes were washed twice for 15 min at room temperature with 2x SSC/0.1% sodium dodecyl sulfate (SDS), followed by two 15-min washes at 48 C with 0.1x SSC/0.1% SDS. After 1 h blocking at room temperature with blocking reagent (Roche), the membranes were incubated with first anti-DIG-alkaline phosphatase conjugate (Roche) and then the chemiluminescent substrate CDP-Star (Roche). Finally, membranes were exposed to Hyperfilm ECL (Amersham Biosciences, Barcelona, Spain). The films were scanned in an Agfa DUOSCAN densitometer, and the bands were quantified using the Kodak 1D Image Analysis Software 3.5 for Windows. Finally, blots were stripped by 10 min exposure to boiling 0.1% SDS and reprobed for 18S rRNA detection to check the loading and transfer of RNA during the blotting. For 18S rRNA, the 31-mer DIG-labeled antisense oligonucleotide 5'-CGCCTGCTGCCTTCCTTGGATGTGGTAGCCG-3' at a concentration of 70 pg/ml was used.
Western blot for UCP1 in BAT
BAT was homogenized at 4 C in 1:5 (wt/vol) of PBS in a Teflon glass homogenizer. The homogenate was centrifuged at 7000 x g for 2 min at 4 C and the supernatant used for total protein and UCP1 analysis. Total protein content was measured by the method of Bradford (29). For UCP1 analysis, 5 μg of total protein were fractionated by SDS-PAGE (10% polyacrylamide) according to Laemmli (30) and electrotransferred onto a nitrocellulose membrane (Bio-Rad Laboratories, Madrid, Spain). After blocking, the membrane was incubated with the primary rabbit polyclonal anti-UCP1 antibody ( Diagnostic, San Antonio, TX), diluted 1:1000, and then with the secondary biotinylated antirabbit IgG antibody conjugated to a streptavidin biotinylated horseradish peroxidase complex (Amersham Biosciences) diluted 1:5000. The immunocomplexes were revealed using an enhanced chemiluminescence detection system (Amersham Biosciences). Membranes were exposed to Hyperfilm enhanced chemiluminescence (Amersham Biosciences). The films were scanned and quantified as described above.
Quantification of leptin levels
Stomach samples were homogenized at 4 C in 1:3 (wt/vol) of PBS [137 mM NaCl, 2.7 mM KCl, and 10 mM PB (pH 7.4)] in a Teflon glass homogenizer. The homogenate was centrifuged at 7000 x g for 2 min at 4 C, and the supernatant was used for leptin quantification.
Leptin concentration in the gastric homogenates, serum, and rat milk was measured with a mouse leptin ELISA kit (R&D Systems, Minneapolis, MN).
Quantification of protein, glucose, and triacylglyceride levels
Protein concentration was measured by the method of Bradford (29). Glucose and triacylglyceride concentration was measured enzymatically using commercial kits and following standard procedures (Roche and R-Biopharm, Darmstadt, Germany, for glucose and Sigma for triacylglycerides).
Immunohistochemistry
The tissues were dehydrated in a graded series of ethanol and embedded in paraffin blocks. Immunohistochemical demonstration of leptin was performed in dewaxed 3-μm sections of tissue according to ABC method (31): 0.3% hydrogen peroxide in methanol for 30 min to block endogenous peroxidase; normal goat serum diluted 1:75 in PBS to reduce nonspecific background staining; polyclonal rabbit antileptin antibody (Santa Cruz Biotechnology, Santa Cruz, CA) 1:200 in PBS overnight at 4 C; biotinylated goat antirabbit IgG (Vector Laboratories, Burlingame, CA) 1:200 for 30 min at room temperature; ABC complex (Vectastain ABC kit; Vector) for 60 min at room temperature; and enzymatic development of peroxidase using Fast 3,3'-diaminobenzidine tablet (Sigma, St. Louis, MO) in water for 3 min in a dark room. Sections were counterstained with hematoxylin and then dehydrated in ethanol, cleared in xylol, and mounted in Eukitt (Kindler, Germany). Negative control was performed by omission of primary antibody.
Statistical analysis
Data are expressed as means ± SEM. One-way ANOVA followed by least significance difference post hoc comparisons was used to assess statistical differences between the groups in studies I and II; two-way ANOVA was also used to assess differences between control and leptin-treated animals at different times in study II; Student’s t test was used to assess statistical differences between control and leptin-treated animals in study III. In all cases, threshold of significance was defined at P < 0.05.
Results
Milk leptin concentration in nursing rats during lactation (study I)
Leptin concentration in maternal milk increased during lactation (Table 1). This pattern was not related with changes in the serum concentration of leptin in nursing rats, which remained unchanged during the lactation period. Triacyl-glyceride and total protein milk concentrations remained stable during the lactation period.
TABLE 1. Milk leptin, serum leptin concentration, milk triacylglyceride, and milk protein concentration in nursing rats from different days of lactation (study I)
Leptin concentration in milk was lower than in serum, although the difference lessened during lactation (from 4.9 times lower on d 7 of lactation to 1.7 times lower on d 21).
Short-term effect of the administration of a single oral dose of leptin in 4-d-old rats (study II)
Four-day-old suckling rats treated with an oral dose of leptin showed, after 1 and 4 h of treatment, a lower stomach content of food (P < 0.05, one-way ANOVA), whereas in the vehicle-treated group, gastric content of food remained stable after 1 h and increased in the fourth hour (P < 0.05, one-way ANOVA) (Fig. 1A). It must be pointed out that pups remained with their mothers after leptin/vehicle administration and had free access to milk.
FIG. 1. Weight of gastric contents (A) and serum (B) and gastric (C) leptin levels in 4-d-old rats treated with an oral dose of leptin or vehicle at time 0, 1 h, and 4 h later (study II). Five different litters were used to perform the experiment, and pups from each litter were randomly assigned into the different groups. The values from each litter are expressed as a percentage of its time 0 group. The mean values at time 0 were: gastric contents 249 ± 45 mg, serum leptin levels 1613 ± 551 pg/ml, gastric leptin levels 547 ± 96 pg/g. L, Effect of leptin treatment; L x T, interaction of leptin treatment and time (two-way ANOVA, P < 0.05). Within each graph, bars not sharing a common letter (a, b, c, d) are significantly different (one-way ANOVA, P < 0.05).
Figure 1B shows that oral leptin administration in 4-d-old rats resulted in higher serum leptin levels, compared with control animals (P < 0.05, two-way ANOVA). Serum leptin levels increased significantly after leptin administration, peaking after 1 h (P < 0.05, one-way ANOVA); after 4 h of treatment, serum leptin levels were still higher than in the vehicle-treated animals. Serum leptin levels did not change in the vehicle-treated control group.
Gastric leptin content was also higher in leptin-treated animals vs. their controls (P < 0.05, two-way ANOVA), with the highest levels after 4 h of leptin administration (Fig. 1C). No changes were observed in the vehicle-treated group. These results on gastric leptin content (determined by ELISA) agree with the immunohistochemistry studies of gastric mucosa (Fig. 2). Immunostaining for leptin appeared to be increased after 4 h of oral leptin administration, compared with time 0. In control animals (time 0), the leptin-positive signal was mainly located at the apex of the superficial epithelial cells, showing milk leptin absorption (Fig. 2A). In leptin-treated animals (Fig. 2B), the leptin-positive signal was more intense than at time 0 and was located at different levels of the gastric glands, suggesting that the leptin supplied is being absorbed by the gastric mucosa.
FIG. 2. Immunostaining for leptin in the gastric mucosa in 4-d-old rats treated with an oral dose of leptin: A, time 0, and B, after 4 h of treatment (study II). Leptin immunoreactivity is mainly located in the apex of the superficial epithelial cells. Four hours after leptin administration, immunoreactivity is stronger than in the time 0 group. Light microscopy, x40.
Effect of chronic administration of oral leptin during the lactation period in neonatal rats (study III)
As shown in Fig. 3A, pups treated with a daily oral dose of leptin during lactation displayed at weaning lower gastric content of food (23% reduction) than control animals (P < 0.05, Student’s t test). However, neither body weight at the end of the study (control: 47.1 ± 0.5 g; leptin: 46.6 ± 0.5 g) nor the size of the fat depots studied (results not shown) was affected by leptin treatment. Serum glucose levels (control: 122 ± 6 mg/dl; leptin: 124 ± 5 mg/dl) and serum triacylglyceride levels (control: 259 ± 44 mg/dl; leptin: 200 ± 23 mg/dl) were not affected by leptin treatment.
FIG. 3. Weight of gastric contents (A), gastric leptin concentration (B), and gastric leptin mRNA levels (C) in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). Leptin mRNA levels were measured by RT-PCR and values expressed relative to ?-actin mRNA levels in arbitrary units (Au). Results are expressed as mean ± SEM (n = 5). *, P < 0.05, control vs. leptin-treated rats (Student’s t test). D, Immunostaining for leptin in the gastric mucosa of 21-d-old rats (light microscopy, x40) in control (D1) and leptin-treated animals (D2). The positive staining for leptin in the superficial epithelium of the mucosa is reduced, compared with 4-d-old animals, and the immunoreactivity for leptin is mainly evident in the basal part of the gastric glands. Leptin-positive signal is weaker in the gastric mucosa of leptin-treated animals.
Oral leptin administration during the whole lactation period resulted in lower gastric leptin levels, compared with control animals (P < 0.05, Student’s t test) (Fig. 3B). Lower leptin immunoreactivity was also shown in the gastric mucosa of leptin-treated pups, compared with control animals, suggesting lower leptin production by the gastric mucosa in these animals (Fig. 3D). No significant leptin staining was found at the apex of the superficial epithelial cells in either of the two groups of animals, but leptin staining was found in the basal part of the gastric glands, indicating that in 21-d-old animals, the source of gastric leptin is mainly endogenous production rather than exogenous leptin absorption. Leptin mRNA expression in the stomach was not significantly affected by leptin treatment (Fig. 3C).
Leptin mRNA levels in the sc inguinal fat depot were significantly lower in leptin-treated animals than control animals (P < 0.05, Student’s t test) (Fig. 4B), but no differences were found in leptin expression in internal (epididymal and retroperitoneal) adipose tissue depots (Fig. 4, C and D). Despite a small decrease, circulating leptin levels did not display significant differences between leptin-treated and control animals (Fig. 4A).
FIG. 4. A, Serum leptin concentration in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation. B, C, and D, Leptin mRNA levels in different WAT depots (inguinal, B; epididymal, C; and retroperitoneal, D) in 21-d old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). Leptin mRNA levels were determined by RT-PCR. The representative results for the different WAT depots are shown in the top panel. In the respective bottom panels, all values of leptin mRNA levels are expressed relative to ?-actin mRNA levels (arbitrary units, Au) as mean ± SEM (n = 5). *, P < 0.05, leptin-treated vs. control animals (Student’s t test).
As shown in Fig. 5, the pups who received a chronic oral dose of leptin during lactation displayed, at the end of the treatment, lower UCP1 mRNA levels and lower UCP1 levels in BAT, compared with the vehicle-treated pups (P < 0.05, Student’s t test). A similar tendency was observed for UCP3 in inguinal WAT (control: 0.83 ± 0.28 Au; leptin: 0.30 ± 0.12 Au), although differences were not significant (P = 0.164 Student’s t test). No changes were found in UCP3 mRNA levels in either BAT or skeletal muscle (data not shown).
FIG. 5. A, UCP1 mRNA expression levels (measured by Northern blotting) in the interscapular BAT in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). UCP1 mRNA levels are expressed relative to 18S rRNA levels. B, UCP1 protein levels (measured by Western blotting) in the interscapular BAT in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation. Representative results are shown in the top panels. In the bottom panels, results represent means ± SEM (n = 5) expressed in arbitrary units (Au). *, P < 0.05, leptin-treated vs. control animals (Student’s t test).
Discussion
Leptin is present in human maternal milk (13) and is secreted by mammary epithelial cells in milk fat globules (32). Here we also show that leptin is present in rat milk at a lower concentration than in the serum, and its levels increase throughout the lactation period. Maternal circulating leptin levels do not display any significant change during the lactation period. Serum leptin concentration has been described to increase during gestation, decline after delivery, and remain stable in lactating rats (33). This pattern is similar to what has been reported in women (34), thus suggesting that the rat may be a useful model for the study of leptin during pregnancy. Although we did not determine leptin production by the epithelial mammary cells, the fact that milk leptin content increased during lactation independently of changes in serum leptin levels suggests that these changes are related to differences in leptin production by the epithelial mammary cells.
Leptin in nursing rats can be transferred via milk to the stomach and afterward into the neonatal rat circulation (13), suggesting that maternal milk leptin may play a regulatory role during development. The transfer of ingested leptin to the bloodstream has been described in 9-d-old rats (13), and leptin immunoreactivity has been found in absorbing milk fat droplets in the gastric mucosa of 2-d-old suckling rats (23); these leptin-positive fat droplets were located mainly in the basal part of the epithelium and also in different levels of the epithelial cells, suggesting a maternal milk-leptin source (23). Present results agree with previously published results showing that the leptin present in milk may be absorbed by the gastrointestinal tract and pass to neonatal blood (13, 28). We have seen that a single oral administration of 4 ng of leptin to 4-d-old rat pups resulted in an acute increase (1-fold) in both gastric and serum leptin levels. The amount of leptin supplied may account for the increase in leptin content in both stomach and serum. Immunohistochemistry also confirms that the increase in the gastric content of leptin was mainly due to exogenous leptin absorption by the epithelium rather than higher leptin production in this tissue.
Leptin present in milk appears to be associated with fat globules (32). It has been postulated that the association of leptin with the fat globules may confer a protective effect against the degradation of leptin by the infant digestive tract (32). Here leptin was administered using water as a vehicle, and the results also suggest an effective absorption by the gastrointestinal tract.
Leptin administered to 4-d-old rat pups had significant effects on the gastric contents of food. After 1 h of leptin administration, pups displayed lower gastric contents than the control group, and this reduction lasted 4 h later. This effect was independent of circadian rhythms of leptin because no changes during the first hour and an increase in the fourth hour occurred in the gastric contents in animals treated with the vehicle. These results suggest that oral intake of leptin plays a role in the short-term control of food intake in neonates.
The effects of gastric leptin on brain stem neuronal activity have been previously reported (21). Furthermore, the long (Ob-Rb) and short (Ob-Ra) forms of leptin receptor are present in the nodose ganglion, which contains the cell bodies of the vagal afferent neurons (35). The vagus nerve plays a major role in the control of short-term food intake by transmitting signals arising from the upper gastrointestinal tract to the nucleus of the solitary tract, which is the primary site of projections of vagal afferent fibers (35). Recently it has been reported that leptin is capable of direct and acute activation of vagal afferent neurons, triggering an acute influx of extracellular calcium in these neurons (36). Thus, orally administrated leptin in lactating rats may interact with its receptor in the vagal nerve and control the size of food intake, probably as a short-term signal. In addition, leptin present in the gastric lumen could pass to the bloodstream, which suggests that leptin may have additional systemic effects, probably by acting on its hypothalamic receptors. Thus, exogenous leptin administration could play a dual role in the control of food intake acting either through the activation of afferent vagal nerves (leptin in the gastric lumen) or centrally (leptin in the systemic circulation). Both pathways could participate in the short-term regulation of food intake in the neonatal rats.
In accordance with the postulated inhibitory effect of leptin on feeding during lactation, we observed that when neonatal rats were chronically treated with leptin during the whole lactation period, the weight of gastric contents was significantly lower than that of control animals. However, leptin-treated animals did not display either a reduction in body weight or the size of fat depots, suggesting further metabolic adaptations able to balance the energy intake. Previous results also showed that exogenous administration (via a stomach tube) in neonatal piglets for 6 d of higher doses of leptin than those used here did not produce significant effects on body weight (37).
Oral administration of leptin during lactation was shown to inhibit gastric leptin production: gastric leptin content in 21-d-old animals was lower in leptin-treated animals than control animals, and immunohistochemistry studies also showed that the gastric mucosa appeared less intensely stained in the leptin-treated group than the control group, indicating lower leptin production by the gastric mucosa. Leptin mRNA expression in the stomach of leptin-treated animals was slightly lower than in control animals. This suggests that orally supplied leptin inhibits gastric leptin production possibly through a feedback mechanism acting on the stomach receptors. This is in agreement with our previous observations showing little gastric leptin production during the first days of lactation but higher production with the suspension of suckling and change to solid diet (23). Unlike what was seen in 4-d-old rats, immunohistochemistry did not show evidence of leptin absorption by the gastric mucosa in 21-d-old animals. This may be due to the fact that leptin treatment finished the day before the animals were killed (d 20 of lactation) or may indicate a decrease in leptin absorption capacity throughout lactation, according to the maturation process of the gastric mucosa.
Leptin mRNA expression in the sc inguinal WAT depot, but not in the retroperitoneal and gonadal depots, was lower in leptin-treated animals, which could be the result of lower food intake. Although young rats store a larger proportion of their fat sc, compared with adult rats (38), leptin mRNA production by this depot is lower than by internal visceral depots (28); thus, its contribution to the overall circulating leptin is relatively low. In fact, the decrease in leptin production by the sc depot is not translated into significant changes in blood leptin levels.
UCPs, in particular the UCP1 in BAT, constitute the molecular basis of physiological adaptive thermogenesis in small mammals (39, 40). Both UCP1 mRNA levels and UCP1 levels in the BAT displayed a significant decrease after leptin treatment during lactation. A similar, but slighter, tendency was also observed for UCP3 expression in the inguinal WAT. A previous report shows that ip injection of leptin to neonatal mice produces minor effects on UCP1 mRNA expression in BAT but increases UCP3 mRNA expression (41). Here UCP1 down-regulation in animals orally supplemented with leptin would probably be the result of lower food intake and thus lower diet-induced thermogenesis rather than a direct effect of oral leptin on BAT. A depressed thermogenic capacity, as suggested by lower UCP1 in BAT, helps to explain the maintenance of body weight despite lower food intake.
Thus, several metabolic adaptations occur in leptin-supplemented animals during lactation (namely lower food intake, lower leptin production in the stomach and sc WAT, and lower thermogenic capacity) without changing body weight gain. This would indicate that measurements of body weight changes may not reflect the amount of food intake during the lactation period, which may have implications when translated into humans. Results also point out that differences in food intake between breast-fed infants (which receive different amounts of leptin depending on the nursing mother) and formula-fed infants (which do not receive leptin at all) cannot be underestimated and raises the question as to whether leptin addition to infant formulas should be considered.
In conclusion, leptin exerts important effects during lactation in a physiologically regulated manner. It is absorbed by the rat stomach during neonatal development triggering down-regulation of endogenous leptin production and regulating food intake, without affecting body weight gain, bearing relation with a balanced lower BAT thermogenesis. It is suggested that leptin supplied by milk or ingested from another source during early life plays an important role in the neonatal control of energy balance and possibly in other functions during development.
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Address all correspondence and requests for reprints to: C. Picó, Department of Biología Fundamental y Ciencias de la Salud, Universitat de les Illes Balears, Cra. Valldemossa Km 7.5, E-07122 Palma de Mallorca, Spain. E-mail: cati.pico@uib.es.
Abstract
Although leptin is a hormone mainly produced by the adipose tissue, it is also produced by the gastric mucosa and the mammary epithelium and is present in maternal milk. The effects of milk leptin on the neonate are not known. The purpose of the investigation was to evaluate the short-term effects of the administration of a single oral dose of leptin on 4-d-old rats as well as the effects of chronic supplementation during the lactation period with a daily oral dose of leptin (equivalent to 5 times the amount of leptin ingested normally from maternal milk during the suckling period) on body weight, the gastric leptin system, gastric food content, and thermogenic capacity. Our results show that the administration of a single oral dose of 4 ng of leptin to 4-d-old rats produces a short-term increase in leptin levels in the stomach and serum and a decrease in the weight of the gastric contents. Pups treated with a daily oral dose of leptin during the whole lactation period showed, at the end of the suckling period, compared with controls, lower gastric contents, lower leptin production by the stomach and the sc adipose tissue, and lower thermogenic capacity in brown adipose tissue. We conclude that oral leptin is absorbed by the immature gastric epithelium of the neonate, and this leptin exerts clear biological effects, down-regulating endogenous leptin production and playing a potential role in the short-term control on food intake during the lactation period.
Introduction
LEPTIN IS AN anorexigenic hormone, mainly produced by the adipose tissue (1), which plays an important role in the central regulation of energy balance. This hormone is released into the circulatory system by the adipose tissue in proportion to the amount of lipid stores (2, 3) and acts at the hypothalamic receptors (4), decreasing food intake and increasing energy expenditure (5, 6). Leptin production in nonadipose tissue such as stomach (7, 8, 9), placenta (10, 11), skeletal muscle (12), and mammary epithelium (13) has also been reported, thus extending the role of leptin to various biological functions (5, 14).
Leptin in the human stomach is located in the lower half of the stomach glands, particularly in the pepsinogen granules of chief cells and in a specific endocrine cell-type (8, 9). There is evidence indicating that feeding stimulates the secretion of leptin by the stomach in humans (8, 9) and rats (7, 15), in which a short food intake stimulus (20 min of refeeding) after 14 h of fasting is capable of practically emptying the leptin stores from within the stomach mucosa (15). In addition, gastric leptin shows diurnal variations influenced by food intake rhythms and changes occurring just before the beginning of the feeding period are opposite to those of the orexigenic hormone ghrelin (16). Pepsinogen secretagogues such as cholecystokinin, gastrin, or secretin also induce gastric leptin release in rats (7).
Leptin receptors are present in the human stomach (17, 18, 19) and mouse gastrointestinal tract (20). Leptin can modulate vagal afferent fibers that originate in the gastric and intestinal walls and terminate in the nucleus tractus solitarius (21), providing rapid information to the brain. These data, together with the observed sensitivity of gastric leptin to the nutritional state, have allowed us to suggest a role for gastric leptin in the short-term regulation of food intake, probably acting as a satiety signal (22).
In neonate rats, leptin production by the stomach probably starts at the onset of suckling, although leptin production by the stomach is kept at low levels during the suckling period (23). The change of diet from milk to solid chow is associated with a significant increase in gastric leptin production (23). The presence of leptin in maternal milk and the fact that leptin in nursing rats could be absorbed by the gastric mucosa of neonatal rats (23) and transferred to the infant rat circulation (13) have suggested that leptin supplied from maternal milk could be the main source of leptin in the stomach during the suckling period (23). This exogenous leptin could exert biological effects in neonates at a time in which both the adipose tissue and appetite regulatory systems are immature (21).
Leptin concentrations in human milk vary significantly between people (24, 25), and some authors have found a positive correlation between leptin concentration in milk and maternal plasma leptin levels and adiposity (24). Thus, breast-fed infants nursed by mothers with significant adiposity may be exposed to higher amounts of leptin than infants nursed by lean mothers, and much higher than those fed with infant formulas, which do not have leptin as an ingredient (26). However, the effects of oral intake of leptin on neonate development are not known.
The aim of this study was to evaluate the effects of the administration of an acute oral dose of leptin on food intake in neonatal rats as well as the prolonged effects of the chronic administration of a daily oral dose of leptin, close to physiological levels, during the suckling period on the gastric leptin system and energy balance to determine the importance of leptin supplied by maternal milk during lactation.
Materials and Methods
Animals and experimental design
Study I: study of milk leptin concentration in nursing rats during lactation.
Three-month-old, virgin female Wistar rats were caged with a male rat. Animals were from Charles River Laboratories (Barcelona, Spain). After matching, each female was placed in an individual cage with free access to water and food. Rats were kept in a room with controlled temperature (22 C) and a 12-h light, 12-h dark cycle (light on from 0800 to 2000 h). After delivery, nine dams were used for milk studies. Birth was defined as d 0 of lactation. Milk and serum samples of each dam were collected on d 7, 14, and 21 of lactation. For milk collection nursing rats were separated from their pups for 6 h to guarantee that mammary glands were full of milk. Before milking, dams were exposed to ether, and then milk was obtained from the mammary glands by manual milking and stored frozen at –20 C. A sample of blood was also collected from the end of the tail, stored at room temperature for 1 h and overnight at 4 C, and then centrifuged at 1000 x g for 10 min to collect the serum.
Study II: short-term effect of the administration of a single oral dose of leptin in 4-d-old rats.
Four nanograms of recombinant murine leptin (PeproTech, London, UK) or vehicle (water) were orally administrated with a micropipette to 4-d-old Wistar suckling rats belonging to five different litters. After leptin/vehicle administration, pups were kept with their mothers. Rats were killed by decapitation at different times: time 0, just before leptin administration, and 1 and 4 h after leptin or vehicle administration. A sample of blood was collected and processed as described above (study I) to obtain the serum. The stomach was immediately removed in its entirety, opened, rinsed with saline containing 0.1% diethyl pyrocarbonate (Sigma, Madrid, Spain), frozen in liquid nitrogen, and stored at –70 C until analysis. A lengthway fragment from the stomach was fixed by immersion in 4% paraformaldehyde in 0.1 M phosphate buffer (PB) (pH 7.4) overnight at 4 C for immunohistochemistry studies. Gastric contents were weighed.
Study III: effect of chronic administration of oral leptin during the lactation period in neonatal rats.
At d 1 after delivery, pups from the same litter were randomly distributed into two groups: control group and leptin-treated group (n = 5 in each group). From d 1 to d 20 of lactation and during the first 2 h of the beginning of the light cycle, 20 μl of the vehicle (water) or a solution of recombinant murine leptin (PeproTech) was given orally every day to the pups using a pipette. The amount of leptin given to animals was progressively increased from 1 ng of leptin on d 1 to 43.8 ng of leptin on d 20. This dose of leptin was calculated as 5 times the amount of the daily leptin intake from maternal milk, which was calculated from leptin concentration in milk during the lactation period (obtained from study I) and the estimated total daily milk intake during the lactation period (27). We considered this dose of leptin as close to physiological levels, taking into account the range of variation of milk leptin levels in dams. At weaning, on d 21, pups were killed and the stomach, gastrocnemius muscle, brown adipose tissue (BAT), and different white adipose tissue depots (WAT) (epididymal, retroperitoneal, and inguinal) were rapidly removed. The stomach was opened and rinsed with saline containing 0.1% diethyl pyrocarbonate (Sigma). The gastric contents and BAT and WAT were weighed. All samples were immediately frozen in liquid nitrogen and stored at –70 C until RNA analysis. A lengthway fragment from the stomach was fixed by immersion in 4% paraformaldehyde in 0.1 M PB (pH 7.4) overnight at 4 C. Blood was also collected and processed as described in study I to obtain the serum.
The animal protocol followed in this study was reviewed and approved by the bioethical committee of our university and guidelines for the use and care of laboratory animals of the university were followed.
RT-PCR analysis of leptin mRNA in the stomach and WAT
To determine the expression of leptin in the stomach and WAT, we developed a RT-PCR using the housekeeping gene ?-actin as internal control. Total RNA was extracted using Trizol (Invitrogen, Barcelona, Spain) reagent according to the instructions provided by the supplier; 0.25 μg of total RNA (in a final volume of 5 μl) was denatured at 90 C for 1 min and then reverse transcribed to cDNA using Moloney leukemia virus reverse transcriptase (according to the procedure of Applied Biosystems, Foster City, CA) at 42 C for 1 h, with a final step of 5 min at 99 C in a 2400 thermal cycler (PerkinElmer, Norwalk, CT). Sample was denatured at 94 C for 3 min, and then PCR was carried out using the following parameters: 94 C for 1 min, 58 C for 1 min, and 72 C for 2 min. Thirty cycles were carried out for stomach samples and 23 cycles for the WAT samples. The amplification was finished by an extension step of 10 min at 72 C. Primers for the lep gene were: forward 5'-CCA GGA TGA CAC CAA AAC CCT C-3' and reverse 5'-ATC CAG GCT CTC TGG CTT CTG C-3', and for the ?-actin gene: forward 5'-ACG GGC ATT GTG ATG GAC TC-3' and reverse 5'-GTG GTG GTG AAG CTG TAG CC-3'. The expected size of the products were 316 bp for the lep gene and 164 bp for the ?-actin gene, which were visualized by electrophoresis in a 1.5% agarose gel containing ethidium bromide and verified by using a DNA 100-bp ladder. The bands in the gel were quantified by scanner photodensitometry using the Kodak 1D Image Analysis software (version 3.5 for Windows; Eastman Kodak Co., Rochester, NY). The signal for leptin mRNA was normalized to the signal of the housekeeping gene ?-actin and the results were expressed as the leptin to ?-actin mRNA ratio.
Northern blot analysis of uncoupling protein (UCP)1 and UCP3 mRNA
mRNA levels for UCP1 in BAT and UCP3 in gastrocnemius muscle, BAT, and inguinal WAT were determined by Northern blot. Thirty micrograms of total RNA were denatured with formamide/formaldehyde and then fractionated by agarose gel electrophoresis as previously described (28). The RNA was transferred onto a nylon membrane (Roche, Barcelona, Spain) in 20x saline sodium citrate (SSC) buffer: 1x SSC is 150 mM NaCl, 15 mM sodium citrate (pH 7.0) by capillary blotting for 16 h and fixed with UV light.
The specific mRNA were detected by a chemiluminescence based procedure, using a 30-mer antisense oligonucleotide probe (5'-GTTGGTTTTATTCGTGGTCTCCCAGCATAG-3') for UCP1 detection and a 32-mer antisense oligonucleotide probe (5'-GACTCCTTCTTCCCTGGCGATGGTTCTGTAGG-3') for UCP3, both probes synthesized commercially (Roche), and labeled at both ends with a single digoxigenin (DIG) ligand. Prehybridization was at 42 C for 15 min in DIG-Easy Hyb (Roche). Hybridization was at 42 C overnight in DIG-Easy Hyb containing 34 ng/ml of the oligonucleotide probe. Then hybridized membranes were washed twice for 15 min at room temperature with 2x SSC/0.1% sodium dodecyl sulfate (SDS), followed by two 15-min washes at 48 C with 0.1x SSC/0.1% SDS. After 1 h blocking at room temperature with blocking reagent (Roche), the membranes were incubated with first anti-DIG-alkaline phosphatase conjugate (Roche) and then the chemiluminescent substrate CDP-Star (Roche). Finally, membranes were exposed to Hyperfilm ECL (Amersham Biosciences, Barcelona, Spain). The films were scanned in an Agfa DUOSCAN densitometer, and the bands were quantified using the Kodak 1D Image Analysis Software 3.5 for Windows. Finally, blots were stripped by 10 min exposure to boiling 0.1% SDS and reprobed for 18S rRNA detection to check the loading and transfer of RNA during the blotting. For 18S rRNA, the 31-mer DIG-labeled antisense oligonucleotide 5'-CGCCTGCTGCCTTCCTTGGATGTGGTAGCCG-3' at a concentration of 70 pg/ml was used.
Western blot for UCP1 in BAT
BAT was homogenized at 4 C in 1:5 (wt/vol) of PBS in a Teflon glass homogenizer. The homogenate was centrifuged at 7000 x g for 2 min at 4 C and the supernatant used for total protein and UCP1 analysis. Total protein content was measured by the method of Bradford (29). For UCP1 analysis, 5 μg of total protein were fractionated by SDS-PAGE (10% polyacrylamide) according to Laemmli (30) and electrotransferred onto a nitrocellulose membrane (Bio-Rad Laboratories, Madrid, Spain). After blocking, the membrane was incubated with the primary rabbit polyclonal anti-UCP1 antibody ( Diagnostic, San Antonio, TX), diluted 1:1000, and then with the secondary biotinylated antirabbit IgG antibody conjugated to a streptavidin biotinylated horseradish peroxidase complex (Amersham Biosciences) diluted 1:5000. The immunocomplexes were revealed using an enhanced chemiluminescence detection system (Amersham Biosciences). Membranes were exposed to Hyperfilm enhanced chemiluminescence (Amersham Biosciences). The films were scanned and quantified as described above.
Quantification of leptin levels
Stomach samples were homogenized at 4 C in 1:3 (wt/vol) of PBS [137 mM NaCl, 2.7 mM KCl, and 10 mM PB (pH 7.4)] in a Teflon glass homogenizer. The homogenate was centrifuged at 7000 x g for 2 min at 4 C, and the supernatant was used for leptin quantification.
Leptin concentration in the gastric homogenates, serum, and rat milk was measured with a mouse leptin ELISA kit (R&D Systems, Minneapolis, MN).
Quantification of protein, glucose, and triacylglyceride levels
Protein concentration was measured by the method of Bradford (29). Glucose and triacylglyceride concentration was measured enzymatically using commercial kits and following standard procedures (Roche and R-Biopharm, Darmstadt, Germany, for glucose and Sigma for triacylglycerides).
Immunohistochemistry
The tissues were dehydrated in a graded series of ethanol and embedded in paraffin blocks. Immunohistochemical demonstration of leptin was performed in dewaxed 3-μm sections of tissue according to ABC method (31): 0.3% hydrogen peroxide in methanol for 30 min to block endogenous peroxidase; normal goat serum diluted 1:75 in PBS to reduce nonspecific background staining; polyclonal rabbit antileptin antibody (Santa Cruz Biotechnology, Santa Cruz, CA) 1:200 in PBS overnight at 4 C; biotinylated goat antirabbit IgG (Vector Laboratories, Burlingame, CA) 1:200 for 30 min at room temperature; ABC complex (Vectastain ABC kit; Vector) for 60 min at room temperature; and enzymatic development of peroxidase using Fast 3,3'-diaminobenzidine tablet (Sigma, St. Louis, MO) in water for 3 min in a dark room. Sections were counterstained with hematoxylin and then dehydrated in ethanol, cleared in xylol, and mounted in Eukitt (Kindler, Germany). Negative control was performed by omission of primary antibody.
Statistical analysis
Data are expressed as means ± SEM. One-way ANOVA followed by least significance difference post hoc comparisons was used to assess statistical differences between the groups in studies I and II; two-way ANOVA was also used to assess differences between control and leptin-treated animals at different times in study II; Student’s t test was used to assess statistical differences between control and leptin-treated animals in study III. In all cases, threshold of significance was defined at P < 0.05.
Results
Milk leptin concentration in nursing rats during lactation (study I)
Leptin concentration in maternal milk increased during lactation (Table 1). This pattern was not related with changes in the serum concentration of leptin in nursing rats, which remained unchanged during the lactation period. Triacyl-glyceride and total protein milk concentrations remained stable during the lactation period.
TABLE 1. Milk leptin, serum leptin concentration, milk triacylglyceride, and milk protein concentration in nursing rats from different days of lactation (study I)
Leptin concentration in milk was lower than in serum, although the difference lessened during lactation (from 4.9 times lower on d 7 of lactation to 1.7 times lower on d 21).
Short-term effect of the administration of a single oral dose of leptin in 4-d-old rats (study II)
Four-day-old suckling rats treated with an oral dose of leptin showed, after 1 and 4 h of treatment, a lower stomach content of food (P < 0.05, one-way ANOVA), whereas in the vehicle-treated group, gastric content of food remained stable after 1 h and increased in the fourth hour (P < 0.05, one-way ANOVA) (Fig. 1A). It must be pointed out that pups remained with their mothers after leptin/vehicle administration and had free access to milk.
FIG. 1. Weight of gastric contents (A) and serum (B) and gastric (C) leptin levels in 4-d-old rats treated with an oral dose of leptin or vehicle at time 0, 1 h, and 4 h later (study II). Five different litters were used to perform the experiment, and pups from each litter were randomly assigned into the different groups. The values from each litter are expressed as a percentage of its time 0 group. The mean values at time 0 were: gastric contents 249 ± 45 mg, serum leptin levels 1613 ± 551 pg/ml, gastric leptin levels 547 ± 96 pg/g. L, Effect of leptin treatment; L x T, interaction of leptin treatment and time (two-way ANOVA, P < 0.05). Within each graph, bars not sharing a common letter (a, b, c, d) are significantly different (one-way ANOVA, P < 0.05).
Figure 1B shows that oral leptin administration in 4-d-old rats resulted in higher serum leptin levels, compared with control animals (P < 0.05, two-way ANOVA). Serum leptin levels increased significantly after leptin administration, peaking after 1 h (P < 0.05, one-way ANOVA); after 4 h of treatment, serum leptin levels were still higher than in the vehicle-treated animals. Serum leptin levels did not change in the vehicle-treated control group.
Gastric leptin content was also higher in leptin-treated animals vs. their controls (P < 0.05, two-way ANOVA), with the highest levels after 4 h of leptin administration (Fig. 1C). No changes were observed in the vehicle-treated group. These results on gastric leptin content (determined by ELISA) agree with the immunohistochemistry studies of gastric mucosa (Fig. 2). Immunostaining for leptin appeared to be increased after 4 h of oral leptin administration, compared with time 0. In control animals (time 0), the leptin-positive signal was mainly located at the apex of the superficial epithelial cells, showing milk leptin absorption (Fig. 2A). In leptin-treated animals (Fig. 2B), the leptin-positive signal was more intense than at time 0 and was located at different levels of the gastric glands, suggesting that the leptin supplied is being absorbed by the gastric mucosa.
FIG. 2. Immunostaining for leptin in the gastric mucosa in 4-d-old rats treated with an oral dose of leptin: A, time 0, and B, after 4 h of treatment (study II). Leptin immunoreactivity is mainly located in the apex of the superficial epithelial cells. Four hours after leptin administration, immunoreactivity is stronger than in the time 0 group. Light microscopy, x40.
Effect of chronic administration of oral leptin during the lactation period in neonatal rats (study III)
As shown in Fig. 3A, pups treated with a daily oral dose of leptin during lactation displayed at weaning lower gastric content of food (23% reduction) than control animals (P < 0.05, Student’s t test). However, neither body weight at the end of the study (control: 47.1 ± 0.5 g; leptin: 46.6 ± 0.5 g) nor the size of the fat depots studied (results not shown) was affected by leptin treatment. Serum glucose levels (control: 122 ± 6 mg/dl; leptin: 124 ± 5 mg/dl) and serum triacylglyceride levels (control: 259 ± 44 mg/dl; leptin: 200 ± 23 mg/dl) were not affected by leptin treatment.
FIG. 3. Weight of gastric contents (A), gastric leptin concentration (B), and gastric leptin mRNA levels (C) in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). Leptin mRNA levels were measured by RT-PCR and values expressed relative to ?-actin mRNA levels in arbitrary units (Au). Results are expressed as mean ± SEM (n = 5). *, P < 0.05, control vs. leptin-treated rats (Student’s t test). D, Immunostaining for leptin in the gastric mucosa of 21-d-old rats (light microscopy, x40) in control (D1) and leptin-treated animals (D2). The positive staining for leptin in the superficial epithelium of the mucosa is reduced, compared with 4-d-old animals, and the immunoreactivity for leptin is mainly evident in the basal part of the gastric glands. Leptin-positive signal is weaker in the gastric mucosa of leptin-treated animals.
Oral leptin administration during the whole lactation period resulted in lower gastric leptin levels, compared with control animals (P < 0.05, Student’s t test) (Fig. 3B). Lower leptin immunoreactivity was also shown in the gastric mucosa of leptin-treated pups, compared with control animals, suggesting lower leptin production by the gastric mucosa in these animals (Fig. 3D). No significant leptin staining was found at the apex of the superficial epithelial cells in either of the two groups of animals, but leptin staining was found in the basal part of the gastric glands, indicating that in 21-d-old animals, the source of gastric leptin is mainly endogenous production rather than exogenous leptin absorption. Leptin mRNA expression in the stomach was not significantly affected by leptin treatment (Fig. 3C).
Leptin mRNA levels in the sc inguinal fat depot were significantly lower in leptin-treated animals than control animals (P < 0.05, Student’s t test) (Fig. 4B), but no differences were found in leptin expression in internal (epididymal and retroperitoneal) adipose tissue depots (Fig. 4, C and D). Despite a small decrease, circulating leptin levels did not display significant differences between leptin-treated and control animals (Fig. 4A).
FIG. 4. A, Serum leptin concentration in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation. B, C, and D, Leptin mRNA levels in different WAT depots (inguinal, B; epididymal, C; and retroperitoneal, D) in 21-d old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). Leptin mRNA levels were determined by RT-PCR. The representative results for the different WAT depots are shown in the top panel. In the respective bottom panels, all values of leptin mRNA levels are expressed relative to ?-actin mRNA levels (arbitrary units, Au) as mean ± SEM (n = 5). *, P < 0.05, leptin-treated vs. control animals (Student’s t test).
As shown in Fig. 5, the pups who received a chronic oral dose of leptin during lactation displayed, at the end of the treatment, lower UCP1 mRNA levels and lower UCP1 levels in BAT, compared with the vehicle-treated pups (P < 0.05, Student’s t test). A similar tendency was observed for UCP3 in inguinal WAT (control: 0.83 ± 0.28 Au; leptin: 0.30 ± 0.12 Au), although differences were not significant (P = 0.164 Student’s t test). No changes were found in UCP3 mRNA levels in either BAT or skeletal muscle (data not shown).
FIG. 5. A, UCP1 mRNA expression levels (measured by Northern blotting) in the interscapular BAT in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation (study III). UCP1 mRNA levels are expressed relative to 18S rRNA levels. B, UCP1 protein levels (measured by Western blotting) in the interscapular BAT in 21-d-old rats that received a daily oral dose of leptin or the vehicle during lactation. Representative results are shown in the top panels. In the bottom panels, results represent means ± SEM (n = 5) expressed in arbitrary units (Au). *, P < 0.05, leptin-treated vs. control animals (Student’s t test).
Discussion
Leptin is present in human maternal milk (13) and is secreted by mammary epithelial cells in milk fat globules (32). Here we also show that leptin is present in rat milk at a lower concentration than in the serum, and its levels increase throughout the lactation period. Maternal circulating leptin levels do not display any significant change during the lactation period. Serum leptin concentration has been described to increase during gestation, decline after delivery, and remain stable in lactating rats (33). This pattern is similar to what has been reported in women (34), thus suggesting that the rat may be a useful model for the study of leptin during pregnancy. Although we did not determine leptin production by the epithelial mammary cells, the fact that milk leptin content increased during lactation independently of changes in serum leptin levels suggests that these changes are related to differences in leptin production by the epithelial mammary cells.
Leptin in nursing rats can be transferred via milk to the stomach and afterward into the neonatal rat circulation (13), suggesting that maternal milk leptin may play a regulatory role during development. The transfer of ingested leptin to the bloodstream has been described in 9-d-old rats (13), and leptin immunoreactivity has been found in absorbing milk fat droplets in the gastric mucosa of 2-d-old suckling rats (23); these leptin-positive fat droplets were located mainly in the basal part of the epithelium and also in different levels of the epithelial cells, suggesting a maternal milk-leptin source (23). Present results agree with previously published results showing that the leptin present in milk may be absorbed by the gastrointestinal tract and pass to neonatal blood (13, 28). We have seen that a single oral administration of 4 ng of leptin to 4-d-old rat pups resulted in an acute increase (1-fold) in both gastric and serum leptin levels. The amount of leptin supplied may account for the increase in leptin content in both stomach and serum. Immunohistochemistry also confirms that the increase in the gastric content of leptin was mainly due to exogenous leptin absorption by the epithelium rather than higher leptin production in this tissue.
Leptin present in milk appears to be associated with fat globules (32). It has been postulated that the association of leptin with the fat globules may confer a protective effect against the degradation of leptin by the infant digestive tract (32). Here leptin was administered using water as a vehicle, and the results also suggest an effective absorption by the gastrointestinal tract.
Leptin administered to 4-d-old rat pups had significant effects on the gastric contents of food. After 1 h of leptin administration, pups displayed lower gastric contents than the control group, and this reduction lasted 4 h later. This effect was independent of circadian rhythms of leptin because no changes during the first hour and an increase in the fourth hour occurred in the gastric contents in animals treated with the vehicle. These results suggest that oral intake of leptin plays a role in the short-term control of food intake in neonates.
The effects of gastric leptin on brain stem neuronal activity have been previously reported (21). Furthermore, the long (Ob-Rb) and short (Ob-Ra) forms of leptin receptor are present in the nodose ganglion, which contains the cell bodies of the vagal afferent neurons (35). The vagus nerve plays a major role in the control of short-term food intake by transmitting signals arising from the upper gastrointestinal tract to the nucleus of the solitary tract, which is the primary site of projections of vagal afferent fibers (35). Recently it has been reported that leptin is capable of direct and acute activation of vagal afferent neurons, triggering an acute influx of extracellular calcium in these neurons (36). Thus, orally administrated leptin in lactating rats may interact with its receptor in the vagal nerve and control the size of food intake, probably as a short-term signal. In addition, leptin present in the gastric lumen could pass to the bloodstream, which suggests that leptin may have additional systemic effects, probably by acting on its hypothalamic receptors. Thus, exogenous leptin administration could play a dual role in the control of food intake acting either through the activation of afferent vagal nerves (leptin in the gastric lumen) or centrally (leptin in the systemic circulation). Both pathways could participate in the short-term regulation of food intake in the neonatal rats.
In accordance with the postulated inhibitory effect of leptin on feeding during lactation, we observed that when neonatal rats were chronically treated with leptin during the whole lactation period, the weight of gastric contents was significantly lower than that of control animals. However, leptin-treated animals did not display either a reduction in body weight or the size of fat depots, suggesting further metabolic adaptations able to balance the energy intake. Previous results also showed that exogenous administration (via a stomach tube) in neonatal piglets for 6 d of higher doses of leptin than those used here did not produce significant effects on body weight (37).
Oral administration of leptin during lactation was shown to inhibit gastric leptin production: gastric leptin content in 21-d-old animals was lower in leptin-treated animals than control animals, and immunohistochemistry studies also showed that the gastric mucosa appeared less intensely stained in the leptin-treated group than the control group, indicating lower leptin production by the gastric mucosa. Leptin mRNA expression in the stomach of leptin-treated animals was slightly lower than in control animals. This suggests that orally supplied leptin inhibits gastric leptin production possibly through a feedback mechanism acting on the stomach receptors. This is in agreement with our previous observations showing little gastric leptin production during the first days of lactation but higher production with the suspension of suckling and change to solid diet (23). Unlike what was seen in 4-d-old rats, immunohistochemistry did not show evidence of leptin absorption by the gastric mucosa in 21-d-old animals. This may be due to the fact that leptin treatment finished the day before the animals were killed (d 20 of lactation) or may indicate a decrease in leptin absorption capacity throughout lactation, according to the maturation process of the gastric mucosa.
Leptin mRNA expression in the sc inguinal WAT depot, but not in the retroperitoneal and gonadal depots, was lower in leptin-treated animals, which could be the result of lower food intake. Although young rats store a larger proportion of their fat sc, compared with adult rats (38), leptin mRNA production by this depot is lower than by internal visceral depots (28); thus, its contribution to the overall circulating leptin is relatively low. In fact, the decrease in leptin production by the sc depot is not translated into significant changes in blood leptin levels.
UCPs, in particular the UCP1 in BAT, constitute the molecular basis of physiological adaptive thermogenesis in small mammals (39, 40). Both UCP1 mRNA levels and UCP1 levels in the BAT displayed a significant decrease after leptin treatment during lactation. A similar, but slighter, tendency was also observed for UCP3 expression in the inguinal WAT. A previous report shows that ip injection of leptin to neonatal mice produces minor effects on UCP1 mRNA expression in BAT but increases UCP3 mRNA expression (41). Here UCP1 down-regulation in animals orally supplemented with leptin would probably be the result of lower food intake and thus lower diet-induced thermogenesis rather than a direct effect of oral leptin on BAT. A depressed thermogenic capacity, as suggested by lower UCP1 in BAT, helps to explain the maintenance of body weight despite lower food intake.
Thus, several metabolic adaptations occur in leptin-supplemented animals during lactation (namely lower food intake, lower leptin production in the stomach and sc WAT, and lower thermogenic capacity) without changing body weight gain. This would indicate that measurements of body weight changes may not reflect the amount of food intake during the lactation period, which may have implications when translated into humans. Results also point out that differences in food intake between breast-fed infants (which receive different amounts of leptin depending on the nursing mother) and formula-fed infants (which do not receive leptin at all) cannot be underestimated and raises the question as to whether leptin addition to infant formulas should be considered.
In conclusion, leptin exerts important effects during lactation in a physiologically regulated manner. It is absorbed by the rat stomach during neonatal development triggering down-regulation of endogenous leptin production and regulating food intake, without affecting body weight gain, bearing relation with a balanced lower BAT thermogenesis. It is suggested that leptin supplied by milk or ingested from another source during early life plays an important role in the neonatal control of energy balance and possibly in other functions during development.
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