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Oxyntomodulin stimulates intestinal glucose uptake in rats
GASTROENTEROLOGY 1997;112:1961–1970
Oxyntomodulin Stimulates Intestinal Glucose Uptake in Rats NATHAN L. COLLIE,* ZHUOCHEN ZHU,* SHANA JORDAN,* and JOSEPH R. REEVE, Jr.‡ *Department of Biological Sciences and Institute for Biotechnology, Texas Tech University, Lubbock, Texas; and ‡Department of Medicine, Center for Ulcer Research and Education: Digestive Diseases Research Center, Veterans Administration Wadsworth Center, School of Medicine, University of California Los Angeles, Los Angeles, California
Background & Aims: Enteroglucagon peptides have long been proposed as mediators of intestinal adaptation, including mucosal growth and nutrient absorptive capacity. The hypothesis that infusions of oxyntomodulin, a bioactive form of enteroglucagon, would stimulate glucose and amino acid uptake was tested and its effects were compared with those of glucagon. Methods: Rats were infused intravenously via minipumps with either saline, rat oxyntomodulin (0.47 nmolrkg01rh01), or glucagon (0.88 nmolrkg01rh01) for 7 days, and plasma hormone levels were measured. At death, intestinal dimensions and brush border uptake of D-glucose and L-proline were measured using an in vitro everted sleeve technique. Results: Plasma enteroglucagon and glucagon levels were increased 4and 12-fold, respectively, but there were no effects on food intake, body weight, or intestinal dimensions. In contrast, oxyntomodulin and glucagon significantly stimulated total intestinal glucose uptake capacity by 44% and 53%, respectively, over controls. Oxyntomodulin most potently enhanced glucose uptake in the ileum (215%), whereas glucagon’s greatest effect was in the jejunum (63%–85%). However, neither peptide affected proline uptake. Conclusions: These results support a new, specific action for oxyntomodulin in intestinal adaptation as a glucose uptake stimulator and confirm glucagon’s role as a regulator of glucose uptake.
O
ne of the remarkable properties of the small intestine is its ability to adjust or ‘‘adapt’’ absorptive function under varying dietary, developmental, environmental, or pathological conditions.1 For glucose and amino acids, numerous studies indicate that the specific brush border transporters for those nutrients appear to be regulated by dietary levels of their transported substrate.2 The correlation between dietary nutrients and nutrient uptake rate at once leads to the reasonable suggestion that luminal nutrients might act as the proximate signal regulating transporter activity in the enterocyte. In fact, there is little or no direct evidence for a ‘‘lacoperon’’ model3 in which nutrients themselves, taken up from the gut lumen or blood, modulate transporter expression at the gene or protein levels. Ferraris and / 5e1d$$0005
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Diamond4 clearly pointed out the striking differences in transporter and enzyme regulation between bacterial and intestinal cells. Instead, nutrient transport in enterocytes likely responds to an as yet poorly defined combination of substrate, neuronal, and endocrine signals. Our focus is on the hormonal role of the regulatory gut peptide oxyntomodulin (OXN) in the adaptive regulation of brush border glucose uptake. OXN is one of four major peptides (glicentin and glucagon-like peptide 1 and 2 are the others) derived posttranslationally from a single proglucagon precursor5–7 and cosecreted by L cells in the distal small intestine and colon.8,9 We chose to study OXN as a possible mediator of intestinal adaptation for three reasons. First, numerous studies link elevations in OXN and glicentin levels (known collectively as enteroglucagon, because both enteric peptides contain the entire sequence of glucagon) to circumstances characterized by enhanced nutrient absorption, such as diabetes,10 lactation,11 cold adaptation,11 and recovery from intestinal resection.11 – 13 Because the gut hypertrophies in these conditions, enteroglucagon has long been considered a potential mucosal growth factor.14 In addition, a well-documented up-regulation of glucose transporters per enterocyte occurs along with mucosal growth in some cases (e.g., diabetes).15 Thus, OXN also might increase the activity of specific nutrient transporters. Second, OXN is released in response to a complex set of signals, including luminal nutrients (glucose and fatty acids),16 – 18 neurotransmitters,19 and peptides.19 – 21 Hence, OXN is in the likely chain of signals leading from dietary nutrients to adaptation. Third, no studies have tested infusions of synthetic rat OXN on nutrient uptake. Substantial evidence exists for only one physiological action of OXN, the inhibition of acid secretion (oxyntic cell modulator), the effect for which OXN was named.22 – 24 The three rationales in the preceding paragraph might equally apply to all four major proglucagon peptides; Abbreviations used in this paper: carb-free, carbohydrate-free; GLI, total glucagon-like immunoreactivity; OXN, oxyntomodulin; SGLT1, Na//glucose cotransporter 1. q 1997 by the American Gastroenterological Association 0016-5085/97/$3.00
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however, we focus on OXN in this study for an additional, practical reason. Our recent chemical characterization and synthesis of rat OXN allowed us to perform in vivo infusion tests.25 Rat OXN differs in sequence from commercially available porcine OXN at position 33 (rat, arginine; pig, lysine)7,25 but is identical to presumptive human OXN predicted from the human preproglucagon gene sequence.26 Thus, our first goal was to infuse rats with a synthetic duplicate of their endogenous OXN peptide and measure changes in mucosal growth and brush border D-glucose uptake (mediated by the Na//glucose cotransporter 1 [SGLT1])27 along the intestine. Our infusions targeted OXN elevations at physiological levels observed in animals undergoing intestinal adaptation.14 We also measured uptake of L-proline, mediated by a different Na/dependent (imino) transporter, to test the nutrient specificity of adaptation. Our second goal was to compare effects of OXN infusions with those of glucagon for two reasons: except for an octapeptide extension at its carboxyl terminus, OXN is identical to glucagon in structure, and older studies28–30 have long suggested that glucagon stimulates nutrient absorption, along with recent experiments indicating that glucagon mediates short-term (minutes to days) upregulation of glucose uptake.31,32 This reasoning would similarly suggest that we examine the effects of glicentin infusions, because it too contains the entire sequence of glucagon. However, glicentin’s size (69 amino acids) makes it impractical to synthesize and it is not commercially available. Hence, our studies were limited to OXN and glucagon. We tested both peptides in rats fed a carbohydrate-free (carb-free) diet to assess whether glucose uptake could be stimulated by hormones in the absence of dietary levels of the transported substrate. The results provide the first evidence for a role of OXN in intestinal adaptation, one distinct from enteroglucagon’s role in intestinal growth postulated 25 years ago,33 and confirm and extend glucagon’s action at physiological plasma levels as a regulator of glucose uptake.
Materials and Methods Materials Chemicals (reagent grade) were purchased from commercial sources except where noted. Porcine pancreatic glucagon (ú95% purity) used for infusions and synthetic rat (human, bovine, and porcine) glucagon for radioimmunoassay (RIA) were from Sigma Chemical Co. (St. Louis, MO). Rat OXN was synthesized in the Peptide Biochemistry Core Facility of the CURE: Digestive Diseases Research Center (Los Angeles) as described previously.25 All radioisotopes were from Dupont NEN (Wilmington, DE).
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Animals Male Sprague–Dawley rats (200–250 g) obtained from Sasco, Inc. (Omaha, NE) initially were fed rodent chow (Teklad Rodent Diet; Harlan Teklad, Madison, WI) for 1 week after arrival and then switched to Custom Karasov Carbohydrate-free Diet (ICN Biomedicals, Aurora, OH) 2 weeks before surgery and throughout infusion treatments (described in the following paragraph). Laboratory chows typically are high in carbohydrate (approximately 48%),34 which elevates intestinal glucose uptake rates.4,35 The carb-free diet served to depress basal rates of glucose transport (see Results). All animals were housed singly under a 12/12-hour light-dark cycle at 227C. They received food and water ad libitum throughout the experiment and were used under a protocol approved by the Texas Tech University Animal Care and Use Committee.
Peptide Infusions A total of 24 rats were divided into two sets of 12 rats (6 matched pairs for OXN and control infusions and 6 pairs for glucagon and its control infusions). The infusion pairs were matched closely for body weight (an important determinant of intestinal mass), bled at same time at each sampling period (see next paragraph), and killed together to minimize potential circadian variations in endogenous peptide levels and in nutrient transport rates. For 7 days, 1 rat in each pair received intravenous infusions of 0.9% saline (controls) and the treated rat received either OXN or glucagon (0.47 or 0.88 nmolrkg01rh01, respectively, dissolved in saline) via osmotic minipumps (1 mL/h, Alzet Model 2001; Alza Science Products, Palo Alto, CA). From preliminary experiments, we chose the OXN dosage to increase plasma values of enteroglucagon to about 200 pmol/L. This value represented physiological levels measured in cold-acclimated rats,11 a treatment known to increase glucose absorption.36 The twofold-higher glucagon dosage was selected to achieve an equimolar increase in plasma glucagon, given that the metabolic clearance rate of glucagon is twice that of OXN in sham-operated rats.37 Two of the 6 glucagon-control pairs were removed from the study because of blocked blood-sampling catheters, yielding a final sample size for all measurements of 4 glucagon-control pairs and 6 OXN-control pairs.
Surgical Procedures and Blood Sampling Rats were anesthetized intramuscularly with a solution (1 mL/kg body wt) of ketamine HCl (100 mg/mL; ParkeDavis, Morris Plains, NJ) and xylazine (20 mg/mL; Miles Laboratories, Shawnee, KA) mixed in a ratio of 87:13, respectively. A sterile minipump prefilled with either 0.9% saline or peptide was implanted subcutaneously (SC) 2.5 cm to the right of the sternum and its attached Silastic catheter (0.025 ID 1 0.037 OD; Dow Corning, Midland, MI) tunneled SC to the left external jugular, where it was inserted and secured with 5-0 silk suture for venous infusion. A blood-sampling catheter was then implanted as described by Beverly et al.38 to monitor hormone levels. Briefly, the right exterior jugular vein was exposed via a midline incision in the neck and a 4-cm Silastic
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catheter segment inserted toward the heart. The catheter was secured, tunneled SC, and exteriorized at the top of the head through an L-shaped piece of 21-gauge steel tubing, fixed in position with dental acrylic cement and stainless steel screws anchored in the skull. Catheters were maintained patent by filling with a 40% polyvinylpyrrolidone solution containing 50 U heparin/mL and capped with a heat-sealed piece of Tygon tubing. Blood was sampled from unrestrained rats by first removing the polyvinylpyrrolidone-heparin solution from the catheters and then attaching a 15–20-cm length of Tygon tubing, filled with sterile, heparinized saline to the stainless steel cannula. Blood samples (1 mL) were withdrawn from the rat and mixed immediately with a 100-mL solution containing the anti-peptidase aprotinin (500 U Trasylol/mL blood; FBA Pharmaceuticals, New York, NY) and 1.2 mg Na2 ethylenediaminetetraacetic acid. Plasma was separated by centrifugation (3000g at 47C) and stored at 0807C. Blood was collected during surgery (just before minipump implantation); on infusion day 2, 4, and 6 after surgery; and at death (day 7), with all samples taken between 1300 and 1800 before lights out to minimize diurnal variations and feeding effects on endogenous hormone levels.
Whole-Animal Measurements and Intestinal Dimensions Food intake was recorded daily and body mass on alternate days throughout the experiment. On infusion day 7, nonfasted rats were anesthetized as described previously and a final blood sample was taken. The peritoneal cavity was opened and the entire small intestine rinsed with cold (approximately 47C) mammalian Ringer’s without glucose (in mmol/L: NaCl, 128; KCl, 4.7; CaCl2 , 2.5; KH2PO4 , 1.2; MgSO4 , 1.2; and NaHCO3 , 20; pH 7.4 at 377C, 290 mOsm). The intestine was excised quickly, stripped of mesentery and fat, flushed again with Ringer’s, and divided into three equal proximal, middle and distal parts. Each part was suspended against a vertical scale with a 5-g weight attached for length measurements. The segments were then everted over a glass rod, and the wet mass of each segment was measured. Four everted ‘‘sleeves’’ 1.5 cm in length were cut off from each end of the three intestinal regions for glucose and proline uptake measurements (two replicate sleeves for each nutrient). Tissues handled during the previous procedures were maintained at 47C by rinsing or immersing in cold Ringer’s. We also estimated the nominal surface area per centimeter, the proportion of intestinal thickness represented by mucosa (% scrapable mucosa), and dry mass per centimeter in each gut region using techniques described by Diamond and Karasov.35
Nutrient Uptake Measurements We measured active D-glucose and total L-proline uptake across the intestinal brush border membrane (not transepithelial fluxes) using the everted intestinal sleeve method of Karasov and Diamond.39 Briefly, everted tissue sleeves were secured by ligatures onto glass rods (diameter, 4–6 mm based on intestinal region), preincubated for 5 minutes in Ringer’s
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gassed with 95% O2 –5% CO2 at 377C, and then incubated by suspending the rod over a stirring bar rotating at 1200 rpm in a test tube filled with Ringer’s. The incubation Ringer’s solution differed from the preincubation solution only in that the former contains either 50-mmol/L unlabeled D-glucose or L-proline and trace amounts of D-[14C(U)]glucose or L-[a14 C(U)]proline as nutrient probes. The 50-mmol/L concentrations were selected because they greatly exceed transporter Michaelis constants (Km ), resulting in maximal uptake rates (Vmax ).40,41 We added L-[1-3H(N)]glucose in glucose incubations to correct simultaneously for D-glucose in the adherent fluid and D-glucose taken up passively. After a 2-minute incubation (when uptake is linear with time),40 glucose uptake tissues were rinsed in ice-cold Ringer’s without glucose for 20 seconds to reduce the adherent fluid correction. For proline incubations, we added [1,2-3H]polyethylene glycol (mol wt, 4000) to correct for proline in the adherent fluid, yielding total (i.e., active plus passive) proline uptake. In this case, the rinse at the end of the 2-minute incubation was omitted because it would introduce error into the adherent fluid correction (L-glucose and D-glucose have the same diffusion coefficient but polyethylene glycol and L-proline do not). Instead, only the tip of the rod was blotted to remove excess fluid after proline incubations. For the uptake measurements of both nutrients, a 1-cm portion of the incubated tissue was then cut off and placed into a liquid scintillation vial, and its wet mass determined. The tissue was then solubilized in 1 mL of Soluene 350 (Packard, Meriden, CT) at 407C overnight. The following day, scintillation cocktail (10 mL, Hionic-Fluor; Packard) was added and radioactivity counted on a dual-label scintillation counter (LS6500; Beckman, Fullerton, CA). Activity in the samples was corrected for variable quenching, converted to disintegrations per minute, and the uptake rates calculated as described by Karasov and Diamond.39 In addition to expressing uptake rates normalized to wet mass in different gut regions, we calculated the total intestinal uptake capacity for each nutrient by determining the product of the mean uptake rate per milligram for each region and the region’s wet mass and then summing these products for all small intestinal regions.
Plasma Hormone Measurements We measured plasma glucagon and enteroglucagon (i.e., glicentin plus OXN) to document the actual concentrations of hormones achieved by the minipump infusions. All plasma samples were first extracted with 4% trifluoroacetic acid and then centrifuged (5000g) for 20 minutes at 47C. The supernatant was passed through a C-18 Sep-Pak cartridge (Waters, Milford, MA) and eluted with 90% acetonitrile in 0.1% trifluoroacetic acid in a final volume of 1 mL. Extracts were dried in a Speed-Vac (Savant, Holbrook, NY), reconstituted in assay buffer (0.05 mol/L phosphate-buffered saline with 2% Protenate [Baxter, Glendale, CA] at pH 7.4), and assayed using the appropriate antibodies. Glucagon extraction recoveries averaged 92%, and plasma values were adjusted accordingly. Plasma glucagon was measured by an RIA em-
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ploying a glucagon-specific antibody directed toward the C terminus of the peptide (antibody O4A, purchased from Dr. R. H. Unger, University of Texas Southwestern Medical Center, Dallas, TX).42 Plasma enteroglucagon levels were measured by a subtraction RIA.43 Briefly, pancreatic glucagon levels are subtracted from the total glucagon-like immunoreactivity (GLI), measured with antibody 4304 (kindly provided by Dr. Jens Holst, University of Copenhagen, Copenhagen, Denmark) that recognizes the three GLI peptides containing the glucagon residues 6–15 (i.e., glucagon, glicentin, and OXN).44 Although the subtraction assay does not distinguish between the large (glicentin) and small (OXN) forms of enteroglucagon, preliminary experiments in OXN-infused rats showed that the enteroglucagon activity was related directly to the OXN infusion dosage (data not shown). In both assays, glucagon serves as standard and label (receptor-grade 125I-glucagon). After a 4-day incubation at 47C, dextran-coated charcoal was used to separate bound from free label before counting both fractions in a GammaTrac 1193 (TM Analytic, Elk Grove Village, IL). Enteroglucagon levels are reported as picomolar glucagon equivalents. Intra-assay and interassay coefficients of variation were as follows: glucagon, 11% { 3% and 14% { 5%; and GLI, 14% { 2% and 16% { 4%.
Statistical Analysis Significant differences in mean values between control and peptide-infused rats were determined using Student’s t test for paired observations. Values of P ° 0.05 for group differences were considered statistically significant.
Figure 1. Plasma hormone levels (means { SEM) in paired rats infused for 7 days with saline or peptides. (A ) Enteroglucagon (glicentin / OXN, expressed as picomolar glucagon equivalents; solid lines ) and glucagon (dashed lines ) levels in OXN-infused (●) or saline-infused (m) rats. *P õ 0.005; n Å 6 OXN-control pairs. (B ) Glucagon levels in glucagon-infused (●) or saline-infused (m) rats. *P õ 0.005; n Å 4 glucagon-control pairs.
Results Plasma Hormone Levels in Infused Rats Plasma hormone levels were measured throughout the infusion period to document peptide concentrations achieved in control (saline-infused) and peptide-infused rats. As seen in Figure 1A, a 7-day infusion of rat OXN (0.47 nmolrkg01rh01) significantly elevated plasma enteroglucagon levels fourfold to a plateau of 149 pmol/L (mean value of days 2–6; P õ 0.005) compared with controls (37 pmol/L, mean for all 7 days). In contrast, plasma glucagon concentrations were unaffected by OXN infusion (see Figure 1A). Rats infused with glucagon (0.88 nmolrkg01rh01) showed a 12-fold elevation in plasma glucagon levels (165 pmol/L, mean of days 2–4; P õ 0.005) compared with essentially flat basal values of 14 pmol/L averaged over the 7-day infusion period (Figure 1B). Enteroglucagon levels were not measured in glucagon-infused rats because of inadequate plasma sampling. Note that the infusion rates chosen for OXN and glucagon resulted in similar molar elevations of the respective infused peptides. For both peptides, the decrease in plasma concentrations on day 7 represents exhaustion of the pump reservoir. / 5e1d$$0005
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Food Intake, Body Weight, and Intestinal Dimensions Neither OXN nor glucagon elicited significant differences in food intake compared with controls. Food intake fell to a minimum 24 hours after pump implantation in all groups and then rapidly recovered such that feeding rates on infusion day 7 were equivalent to presurgical values (Figure 2A and B). Consistent with the food intake data, there were no significant differences in body weight between control and peptide-infused rats (Figure 2A and B). We measured intestinal dimensions using multiple types of whole-intestinal and regional tissue techniques to test for effects on intestinal growth. Neither peptide significantly affected whole-intestinal dimensions (Figure 3A and B) nor regional gut mass per centimeter (Figure 4A and B). Other measures of regional tissue dimensions (nominal surface area per centimeter, percent scrapable mucosa, and dry mass per centimeter) similarly showed no changes in response to peptide treatments (data not shown). WBS-Gastro
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Effects on Nutrient Uptake Differences in nutrient uptake per centimeter intestine might result from nonspecific alterations in the number of enterocytes per centimeter (reflected in the milligram wet mass per centimeter value) or from cellspecific changes in transport rate per enterocyte (reflected in the uptake rate per milligram of tissue). Because there were no differences in wet mass per centimeter along the gut (Figure 4A and B), uptake rates were normalized per milligram wet mass. As an additional check for the nutrient specificity of the response, we measured the uptake of two nutrients D-glucose and L-proline, both transported across the brush border primarily by Na/-dependent nutrient transporters.4 Finally, we describe the total uptake capacity for each nutrient to assess the integrated contributions of different gut regions to the whole-intestinal response to OXN and glucagon.
sively in a proximal-distal pattern (Figure 5A). Compared with control values, uptake per milligram was stimulated 12% in the duodenum (P ú 0.05), 30%– 40% in the jejunum (P ú 0.05), and 215% in the ileum (P õ 0.001). Glucagon-infused rats also showed elevated glucose uptake per milligram, but the regional response was distinct from that of OXN-infused rats (Figure 5B). Uptake was increased strongly over control values by 67% (P õ 0.01) in the duodenum, 63%–85% in the jejunum (P õ 0.01), but only 41% in the ileum (P ú 0.05). In contrast to glucose transport, there was no effect of either peptide on proline uptake per milligram (Figure 6A and B). Small Intestinal Nutrient Uptake Capacities
The fourfold elevation in plasma OXN levels achieved after OXN infusions stimulated active glucose uptake, with the strength of response increasing progres-
Figure 7 shows the effects of peptide infusions on the total intestinal uptake capacity for glucose and proline. Saline-control rats on the carb-free diet had glucose uptake capacities of approximately 16 mmol/day. OXN and glucagon infusions significantly increased the uptake capacity for glucose by similar increments over controls (44% and 53%, respectively) to levels of 24 mmolr
Figure 2. Body weight (BWT) and food intake (FI) in (A ) OXN- or (B ) glucagon-infused rats compared with paired saline-infused controls, described in Figure 1. There were no significant effects of either peptide on food intake or body weight compared with saline controls. (A and B ) BWT: n, saline-infused; m, peptide-infused. FI: s, saline; ●, peptide. Mean values { SEM.
Figure 3. Dimensions of the small intestine (pylorus to ileocecal junctions) in 7-day infused rats at death (Figure 1). There were no significant effects of either (A ) OXN or (B ) glucagon on gut length or wet mass. (A and B ) h, saline-infused; , peptide-infused. Mean values { SEM.
Gut Regional Response to Peptide Infusions
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day01rrat01 (Figure 7A). For comparison, consider that rats adapted to a chow (high carbohydrate) diet for ú2 weeks show glucose uptake capacities of approximately 30 mmol/day (unpublished observations, Collie, March 1996). Hence, OXN and glucagon infusions provide carb-free–adapted animals with about 80% of the glucose uptake capacity of chow rats. Without peptide infusions, however, carb-free rats have only 53% of the glucose uptake capacity of chow rats. The increase in uptake capacity of peptide-infused rats appears specific for glucose because proline uptake capacity was unaffected by the peptide infusions (Figure 7B).
Discussion When we elevated plasma concentrations of OXN or glucagon to levels previously observed in other gut adaptation models, glucose uptake across the brush border was significantly stimulated in the absence of dietary carbohydrate. The effect was specific for glucose because (1) proline uptake was unaffected and (2) no changes in intestinal dimensions were detected, which would have led to increased transport of both nutrients in parallel.
Figure 4. Small intestinal wet mass of 1-cm everted sleeves used in the nutrient uptake measurements from duodenum (Duod), jejunum (Jejun), and ileum in infused rats. There were no regional effects of OXN (A, ) or glucagon (B, ) on wet mass per centimeter compared with saline control values (h). For each rat, the value in a given region is the average of four sleeves (two each from the proximal and distal ends of each region), with the means of those averaged values ({ SEM) shown for the 6 OXN-control pairs and 4 glucagon-control pairs.
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The gut regional response to the two peptides was different, with OXN promoting uptake most strongly in the ileum and glucagon, in the proximal two thirds of the intestine. Our results suggest that OXN plays a role in intestinal adaptation not as a growth factor but as a stimulator of active glucose uptake. Enteroglucagon and Glucagon Plasma Levels and Responses Our infusions increased OXN and glucagon to approximately equal plasma levels (149 and 165 pmol/ L, respectively). Plasma enteroglucagon levels reported for different intestinal adaptation models bracket our OXN-infused rat values (in pmol/L): fasted, 3345; fastedrefed, 6845; cold-acclimated/hyperphagic, 20011; and, massive small-bowel resected, 500.12 Similarly, our glucagon-infused rat values are found within the physiological range of plasma glucagon values measured in rats
Figure 5. Active D-glucose uptake across the brush border (normalized per milligram wet mass) in different small intestinal regions of 7-day–infused rats. Uptake was measured in everted sleeves for 2 minutes at 50 mmol/L luminal D-glucose, when influx was linear with time. Values are corrected for both passive uptake and adherent fluid. , Peptide-infused rats; h, saline-infused rats. (A ) OXN infusion increased D-glucose uptake in all regions, but the values were statistically significant only in the ileum (**P õ 0.001; n Å 6 OXN-control pairs). (B ) Glucagon infusion also stimulated D-glucose uptake significantly in all regions (*P õ 0.01, n Å 4 glucagon-control pairs), except in the ileum. Mean value { SEM, with the regional value determined from two adjacent sleeves in each region per rat.
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under various conditions (in pmol/L): resting, 29–5746,47; 6-hour cold exposure, 6648; insulin treatment, 11046; intense exercise, 17946; and fasted neonatal, 146–301.49 Thus, the increase in glucose uptake observed in this study was obtained at physiological OXN and glucagon levels. Debnam and Sharp31 showed recently that chronic glucagon injections (50 mg given intraperitoneally three times daily for 3 days) stimulated galactose uptake in isolated, upper, and middle villus jejunal cells by 31% and 74%, respectively, through a phlorizin-sensitive (i.e., SGLT1mediated) pathway. Because glucose and galactose are both transported actively by SGLT1, their data agree closely with our 63%–85% stimulation by glucagon of glucose uptake in intact jejunal tissue. However, it should be pointed out that their dose, used in several earlier studies,28,29,50,51 results in supraphysiological glucagon elevations (approximately 1800 pg/mL or 516 pmol/L).50 OXN and Glucagon Interactions It is clear that the effects of OXN on glucose uptake were not mediated by glucagon, because glucagon
Figure 6. Total L-proline uptake across the brush border (normalized per milligram wet mass) in different small intestinal regions of 7-day– infused rats. Uptake was measured as described for D-glucose (Figure 5) except that uptake is uncorrected for passive transport (see Materials and Methods for details). Unlike D-glucose, L-proline uptake was unaffected by either OXN infusion (A, ) or glucagon infusion (B, ) compared with control values (h). Values and group size are as described in Figure 5.
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levels were not increased in OXN-infused rats (Figure 1A). However, because we did not measure enteroglucagon levels in glucagon-infused rats, the possibility exists that glucagon’s effects resulted from OXN release in response to glucagon infusion. Four observations argue against this possibility. First, the gut regional responsiveness is different for the two peptides (Figure 5), with OXN most effective in the ileum and glucagon in the duodenum and jejunum. Second, previous studies show no effect of glucagon on enteroglucagon or OXN release by the intestine.20,37 Hence, it is unlikely that OXN levels were elevated in our glucagon-infused rats. Third, in most cases the conditions and signals for glucagon and OXN release are quite different. For example, glucagon is released during starvation, exercise, and hypoglycemia,52 whereas OXN is released after a meal and in hyperphagic conditions. The two exceptions are in diabetes 30,52,53 and cold acclimation,48 when both peptides are elevated. In the latter conditions, fuels must be mobilized not only to meet pathological (diabetic glucopenia) or thermoregulatory (cold acclimation) metabolic requirements, but also replenished through increased food intake and absorption. Fourth, glucagon appears capable of directly stimulating glucose uptake in isolated enterocytes.31,32 The rapid time course of this response (within 15 min-
Figure 7. Total intestinal uptake capacity for D-glucose and L-proline in peptide-infused rats. Uptake capacity was calculated by summing the uptake for each region (the mean uptake per milligram multiplied by the milligram of tissue per region) along the small intestine’s length. Both OXN (A, ) and glucagon (B, ) significantly stimulated D-glucose uptake capacity. Note that L-proline uptake capacity was not affected by either peptide. *P õ 0.05; **P õ 0.01.
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utes) supports a direct action of glucagon on enterocytes (discussed in the next paragraph). Thus, OXN and glucagon may play some independent and some complementary roles in enhancing glucose uptake. What might be the physiological significance of OXN’s actions in the ileum, an intestinal region that normally has low rates of glucose absorption?4 Under normal conditions, glucose uptake in proximal gut regions is sufficient to handle ingested glucose loads and OXN levels are not chronically elevated to high levels. However, in circumstances where uptake capacity in the proximal intestine is exceeded (e.g., in hyperphagic conditions, diabetes, intestinal resection, and lactation), glucose uptake in the ileum may become important (additional absorptive capacity is required). It is under these same conditions that OXN levels are elevated, acting as a possible signal for transport up-regulation. By continuous peptide infusions, we tried to reproduce the chronic elevations in OXN associated with such conditions. The resulting increased glucose uptake suggests that OXN is capable of increasing glucose uptake capacity, primarily by stimulating uptake in the distal gut. Mechanisms of OXN and Glucagon Actions on Glucose Uptake The everted sleeve technique used in this study measures active glucose uptake39 across the brush border membrane, a process mediated by the transporter SGLT1.54 How might OXN or glucagon stimulate SGLT1 transport of glucose? Both indirect and direct mechanisms are feasible. One or both peptides might act indirectly by releasing other peptides or chemical mediators that directly stimulate glucose uptake. For example, both OXN and glucagon stimulate release of insulin,55 which has been reported recently to exert short-term up-regulation of SGLT1-mediated uptake.56 However, OXN in particular is a weak insulin releaser55 and is considered physiologically insignificant compared with other intestinal incretins7 (e.g., glucagon-like peptide 1 and gastric inhibitory peptide). Thus, although insulin cannot be ruled out as a mediator for glucagon-induced uptake stimulation, it is unlikely to mediate OXN’s effects at the dosages used in this study. A second example of indirect signals might involve hyperglycemia, because glucagon and OXN (at sufficiently high doses) elevate blood glucose levels.55 Hyperglycemia per se is thought to signal increased glucose absorption by an unknown mechanism.2,41 Although we did not measure blood glucose levels in this study, infusions of OXN or glucagon at higher dosages (up to 3 nmolrkg01rh01) did not chronically result in blood glu/ 5e1d$$0005
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cose values above 100 mg/dL (unpublished observations, Collie and Mahaney, 1996). Thus, hyperglycemia probably did not signal the enhanced glucose uptake at the lower dosages described in this report. Direct mechanisms of action on enterocytes themselves would potentially include adenosine 3*,5*-cyclic monophosphate (cAMP) elevations mediated through separate OXN and glucagon receptors. Isolated rat jejunal enterocytes incubated with glucagon (10 nmol/L) show increased glucose uptake within 15 minutes.31 Glucagon incubation (0.1–100 nmol/L) caused dose-dependent increases in intracellular cAMP concentration of these cells over a similar time course.32 In the latter study, raising cAMP levels by adding dibutyryl cAMP and theophylline similarly stimulated glucose uptake. These findings suggest a direct action of glucagon on enterocytes, one mediated by increased cAMP. Although receptor binding by OXN also leads to cAMP elevations, the OXN receptor is probably distinct from that for glucagon, as indicated by differing receptor affinities for the two peptides in different target tissues. OXN is 20-fold more potent than glucagon in elevating cAMP levels in isolated rat gastric glands57 but binds to hepatic glucagon receptors with a 50-fold lower affinity than glucagon.55 Our finding that OXN has a regional profile of glucose uptake stimulation distinct from glucagon’s is consistent with the hypothesis of separate receptors. Along with the recent cloning of the glucagon receptor, Jelinek et al.58 showed that glucagon binding causes intracellular elevations not only in cAMP but also in Ca2/. Both second messengers might influence glucose uptake in multiple ways, by stimulating SGLT1 transcription, stabilizing its transcript, or modulating its posttranslational activity. In this regard, the observations that SGLT1 messenger RNA expression is up-regulated by cAMP59 but negatively regulated by protein kinase C activation60 in LLC-PK1 (porcine kidney) cells are particularly interesting. Recent work in diabetic rats, an animal model with elevated enteroglucagon and glucagon plasma levels, indicates that several transporters involved in intestinal sugar absorption (SGLT1 and facilitative glucose transporter 1 and 5) show increased expression at the messenger RNA and protein levels.15,61,62 One additional mechanism proposed for glucagon’s stimulation of Na/-dependent glucose and amino acid uptake is an increased electrochemical gradient for Na/.32,51 However, our data do not support this hypothesis. An Na/ gradient enhanced by either glucagon or OXN would lead to increased uptake of both glucose and proline; yet we observed an increase in glucose uptake alone. Thus, our findings are consistent with the specific induction of SGLT1 by OXN and glucagon. WBS-Gastro
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Future Work Currently, we are testing a wide range of OXN and glucagon doses to establish a full dose-response curve for their effects on glucose uptake. Although we compared the effects of the two peptides in this report at approximately the same plasma levels, the glucagon elevation represents a much larger relative increase (12-fold) than that for OXN (4-fold) over basal values. It is possible that at different concentrations the gut regional pattern of response may be altered or mucosal growth stimulated. However, at the OXN dosages infused, we found no evidence for the mucosal growth effect predicted from previous correlations of gut hypertrophy and enteroglucagon plasma levels. Recently, Gregor et al.63 summarized findings that suggest enteroglucagon may even inhibit enterocyte proliferation. Thus, the connection between this peptide family and mucosal growth regulation remains elusive. Finally, the mechanism of OXN and glucagon action on glucose transport, whether direct or indirect, is currently unclear. The availability of molecular probes for transporter expression and of better cell culture systems, combined with in vivo studies, will help fill in the missing steps that must ultimately link food intake, signal release, and intestinal adaptation.
References 1. Karasov WH, Diamond JM. Adaptive regulation of sugar and amino acid transport by vertebrate intestine. Am J Physiol 1983; 245:G443–G462. 2. Ferraris RP, Diamond JM. Regulation of intestinal nutrient transport. In: Johnson LR, ed. Physiology of the gastrointestinal tract. New York: Raven, 1994:1821–1844. 3. Jacob F, Monod J. Genetic regulatory mechanisms in the synthesis of protein. J Mol Biol 1961;3:318–356. 4. Ferraris RP, Diamond JM. Specific regulation of intestinal nutrient transporters by their dietary substrates. Annu Rev Physiol 1989; 51:125–141. 5. Heinrich G, Gros P, Habener JF. Glucagon gene sequence. J Biol Chem 1984;259:14082–14087. 6. Mojsov S, Heinrich G, Wilson IB, Ravazzola M, Orci L, Habener JF. Preproglucagon gene expression in pancreas and intestine diversifies at the level of post-translational processing. J Biol Chem 1986;261:11880–11889. 7. Holst JJ, Ørskov C. Glucagon and other proglucagon-derived peptides. In: Walsh JH, Dockray GJ, eds. Gut peptides: biochemistry and physiology. New York: Raven, 1994:305–340. 8. Bloom SR, Polak JM. Enteroglucagon and the gut hormone profile of intestinal adaptation. In: Robinson JWL, Dowling RH, Riecken EO, eds. Mechanisms of intestinal adaptation. 1st ed. Lancaster, PA: MTP, 1982:189–198. 9. Jin SLC, Hynes MA, Lund PK. Ontogeny of glucagon messenger RNA and encoded precursor in the rat intestine. Regul Pept 1990; 29:117–131. 10. Heding LG, Rasmussen SM. Determination of pancreatic and gut glucagon-like immunoreactivity (GLI) in normal and diabetic subjects. Diabetologia 1972;8:408–411. 11. Jacobs LR, Bloom SR, Dowling H. Response of plasma and tissue levels of enteroglucagon immunoreactivity to intestinal resection, lactation and hyperphagia. Life Sci 1981;29:2003–2007.
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12. Gornacz GE, Ghatei MA, Al-Mukhtar MYT, Yeats JC, Adrian TE, Wright NA, Bloom SR. Plasma enteroglucagon and CCK levels and cell proliferation in defunctioned small bowel in the rat. Dig Dis Sci 1984;29:1041–1049. 13. Taylor RG, Beveridge DJ, Fuller PJ. Expression of ileal glucagon and peptide tyrosine-tyrosine genes: response to inhibition of polyamine synthesis in the presence of massive small-bowel resection. Biochem J 1992;286:737–741. 14. Bloom SR, Polak JM. The hormonal pattern of intestinal adaptation. Scand J Gastroenterol 1982;72:93–103. 15. Burant CF, Flink S, Depaoli AM, et al. Small intestine hexose transport in experimental diabetes: increased transporter mRNA and protein expression in enterocytes. J Clin Invest 1994;93: 578–585. 16. Unger RH, Ohneda A, Valberde I, Eisentraut AM, Exton J. Characterization of the response of circulating glucagon-like immunoreactivity to intraduodenal and intravenous administration of glucose. J Clin Invest 1968;47:48–65. 17. Jian R, Besterman HS, Sarson DL, Aymes C, Hostein J, Bloom SR, Ramboud JC. Colonic inhibition of gastric secretion in man. Dig Dis Sci 1981;26:195–201. 18. Miazza BM, Al-Mukhtar MYT, Salmeron M, Ghatei MA, Felce-Dachez M, Filali A, Villet R, Wright NA, Bloom SR. Hyperenteroglucagonaemia and small intestinal mucosal growth after colonic perfusion of glucose in rats. Gut 1985;26:518–524. 19. Buchan AMJ, Barber DL, Gregor M, Soll AH. Morphologic and physiologic studies of canine ileal enteroglucagon-containing cells in short-term culture. Gastroenterology 1987;93:791–800. 20. Brubaker PL. Regulation of intestinal proglucagon-derived peptide secretion by intestinal regulatory peptides. Endocrinology 1991; 128:3175–3182. 21. Roberge JN, Gronau KA, Brubaker PL. Gastrin-releasing peptide is a novel mediator of proximal nutrient-induced proglucagonderived peptide secretion from the distal gut. Endocrinology 1996;137:2383–2388. 22. Dubrasquet JM, Bataille D, Gespach C. Oxyntomodulin (glucagon37 or bioactive enteroglucagon): a potent inhibitor of pentagastrin-stimulated acid secretion in rats. Biosci Rep 1982;2:391– 395. 23. Jarrousse C, Audousset-Puech MP, Dubrasquet M, Niel H, Martinez J, Bataille D. Oxyntomodulin (glucagon 37) and its C-terminal octapeptide inhibit gastric acid secretion. FEBS Lett 1985;188: 81–84. 24. Le Quellec A, Kervran A, Blache P, Ciurana AJ, Bataille D. Oxyntomodulin-like immunoreactivity: diurnal profile of a new potential enterogastrone. J Clin Endocrinol Metab 1992;74:1405–1409. 25. Collie NL, Walsh JH, Wong HC, Shively JE, Davis MT, Lee TD, Reeve JR Jr. Purification and sequence of rat oxyntomodulin. Proc Natl Acad Sci USA 1994;91:9362–9366. 26. Bell GI, Sanchez-Pescador R, Laybourn PJ, Najarian RC. Exon duplication and divergence in the human preproglucagon gene. Nature 1983;304:368–371. 27. Hediger MA, Rhoads DB. Molecular physiology of sodium-glucose cotransporters. Physiol Rev 1994;74:993–1026. 28. Rudo ND, Rosenberg IH. Chronic glucagon administration enhances intestinal transport in the rat. Proc Soc Exp Biol Med 1973;142:521–525. 29. Debnam ES. Effect of sodium concentration and plasma sugar concentration on hexose absorption by the rat jejunum in vivo. Pflu¨gers Arch 1982;393:104–108. 30. Riecken EO, Gregor M. Glucagon and small-bowel mucosa. Scand J Gastroenterol 1985;20:30–40. 31. Debnam ES, Sharp PA. Acute and chronic effects of pancreatic glucagon on sugar transport across the brush-border and basolateral membranes of rat jejunal enterocytes. Exp Physiol 1993; 78:197–207. 32. Sharp PA, Debnam ES. The role of cyclic AMP in the control of
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sugar transport across the brush-border and basolateral membranes of rat jejunal enterocytes. Exp Physiol 1994;79:203– 214. Gleeson MH, Bloom SR, Polak JM, Henry K, Dowling RH. Endocrine tumour in kidney affecting small bowel structure, motility and absorptive function. Gut 1971;12:773–782. Ferraris RP, Yasharpour S, Lloyd KCK, Mirzayan R, Diamond JM. Luminal glucose concentrations in the gut under normal conditions. Am J Physiol 1990;259:G822–G837. Diamond JM, Karasov WH. Effect of dietary carbohydrate on monosaccharide uptake by mouse small intestine in vitro. J Physiol 1984;349:419–440. Toloza EM, Lam M, Diamond J. Nutrient extraction by cold-exposed mice: a test of digestive safety margins. Am J Physiol 1991;261:G608–G620. Kervran A, Dubrasquet M, Blache P, Martinez J, Bataille D. Metabolic clearance rates of oxyntomodulin and glucagon in the rat: contribution of the kidney. Regul Pept 1990;31:41–52. Beverly JL, Beverly MF, Meguid MM. Alterations in extracellular GABA in the ventral hypothalamus of rats in response to acute glucoprivation. Am J Physiol 1995;269:R1174–R1178. Karasov WH, Diamond JM. A simple method for measuring intestinal solute uptake in vitro. J Comp Physiol 1983;152:105–116. Toloza EM, Diamond J. Ontogenetic development of nutrient transporters in rat intestine. Am J Physiol 1992;263:G593– G604. Karasov WH, Debnam ES. Rapid adaptation of intestinal glucose transport: a brush-border or basolateral phenomenon? Am J Physiol 1987;253:G54–G61. Faloona GR, Unger RH. Glucagon. In: Jaffe BM, Behrman HR, eds. Methods of hormone radioimmunoassay. New York: Academic, 1974:317–330. Adrian TE. Radioimmunoassay. In: Bloom SR, Long RG, eds. Radioimmunoassay of gut regulatory peptides. New York: Praeger, 1982:3–10. Holst JJ, Christiansen J, Ku¨hl C. The enteroglucagon response to intrajejunal infusion of glucose, triglycerides, and sodium chloride, and its relation to jejunal inhibition of gastric acid secretion in man. Scand J Gastroenterol 1976;11:297–304. Kervran A, Blache P, Bataille D. Distribution of oxyntomodulin and glucagon in the gastrointestinal tract and the plasma of the rat. Endocrinology 1987;121:704–713. Harvey WD, Faloona GR, Unger RH. The effect of adrenergic blockade on exercise-induced hyperglucagonemia. Endocrinology 1974;94:1254–1258. Billington CJ, Briggs JE, Link JG, Levine AS. Glucagon in physiological concentrations stimulates brown fat thermogenesis in vivo. Am J Physiol 1991;261:R501–R507. Seitz HJ, Krone W, Wilke H, Tarnowski W. Rapid rise in plasma glucagon induced by acute cold exposure in man and rat. Pflu¨gers Arch 1981;389:115–120. Girard JR, Cuendet GS, Marliss EB, Kervran A, Rieutort M, Assan R. Fuels, hormones, and liver metabolism at term and during the early postnatal period in the rat. J Clin Invest 1973;52:3190– 3200. Rudo ND, Rosenberg IH, Wissler RW. The effect of partial starvation and glucagon treatment on intestinal villus morphology and cell migration. Proc Soc Exp Biol Med 1976;152:277–280. Thompson CS, Debnam ES. Hyperglucagonaemia: effects on ac-
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tive nutrient uptake by the rat jejunum. J Endocrinol 1986;111: 37–42. Unger RH, Orci L. Glucagon secretion, alpha cell metabolism, and glucagon action. In: DeGroot LJ, ed. Endocrinology. New York: Saunders 1995:1337–1353. Rudo ND, Lawrence AM, Rosenberg IH. Treatment with glucagonbinding antibodies alters the intestinal response to starvation in the rat. Gastroenterology 1975;69:1265–1268. Wright EM. The intestinal Na//glucose cotransporter. Annu Rev Physiol 1993;55:575–589. Baldissera FGA, Holst JJ, Knuhtsen S, Hilsted L, Nielsen OV. Oxyntomodulin [glicentin-(33-69)]: pharmacokinetics, binding to liver cell membranes, effects on isolated perfused pig pancreas, and secretion from isolated perfused lower small intestine of pigs. Regul Pept 1988;21:151–166. Stumpel F, Kueera T, Jungermann K. Acute increase by portal insulin in intestinal glucose absorption via hepatoenteral nerves in the rat. Gastroenterology 1996;110:1863–1869. Bataille D, Gespach C, Tatemoto K, Marie J, Coudray AM, Rosselin G, Mutt V. Bioactive enteroglucagon (oxyntomodulin): present knowledge on its chemical structure and its biological activities. Peptides 1981;2:41–44. Jelinek LJ, Lok S, Rosenberg GB, Smith RA, Grant FJ, Biggs S, Bensch P, Kuijper JL, Sheppard PO, Sprecher CA, O’Hara PJ, Foster D, Walker KM, Chen LHJ, McKernan PA, Kindsvogel W. Expression cloning and signaling properties of the rat glucagon receptor. Science 1993;259:1614–1616. Yet SF, Kong CT, Peng H, Lever J. Regulation of Na//glucose cotransporter (SGLT1) mRNA in LLC-PK1 Cells. J Cell Physiol 1994;158:506–512. Shioda T, Ohta T, Isselbacher KJ, Rhoads DB. Differentiationdependent expression of the Na//glucose cotransporter (SGLT1) in LLC-PK1 cells: role of protein kinase C activation and ongoing transcription. Proc Natl Acad Sci USA 1994;91:11919–11923. Burant CF, Saxena M. Rapid reversible substrate regulation of fructose transporter expression in rat small intestine and kidney. Am J Physiol 1994;267:G71–G79. Debnam ES, Smith MW, Sharp PA, Srai SKS, Turvey A, Keable SJ. The effects of streptozotocin diabetes on sodium-glucose transporter (SGLT1) expression and function in rat jejunal and ileal villus-attached enterocytes. Eur J Physiol 1995;430:151– 159. Gregor M, Stallmach A, Menge H, Riecken EO. The role of gutglucagon-like immunoreactants in the control of gastrointestinal epithelial cell renewal. Digestion 1990;46:59–65.
Received August 8, 1996. Accepted January 31, 1997. Address requests for reprints to: Nathan L. Collie, Ph.D., Department of Biological Sciences, Box 43131, Texas Tech University, Lubbock, Texas 79409-3131. Fax: (806) 742-2963. Supported by National Institutes of Health grant DK 42973, a TTU Institute for Nutritional Sciences grant, and a Howard Hughes Medical Institute Fellowship (to S.J.) through the Undergraduate Biological Sciences Education Program. Dr. Zhu’s current affiliation is: Division of Medicine, M. D. Anderson Cancer Center, University of Texas, Houston, Texas 77030. The authors thank Jared Diamond and John Walsh for their support and discussions at critical stages in this project. Wade Mahaney, Richard Lombardini, and Carol Zhang provided valuable assistance in manuscript preparation and graphics.
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Oxyntomodulin Stimulates Intestinal Glucose Uptake in Rats NATHAN L. COLLIE,* ZHUOCHEN ZHU,* SHANA JORDAN,* and JOSEPH R. REEVE, Jr.‡ *Department of Biological Sciences and Institute for Biotechnology, Texas Tech University, Lubbock, Texas; and ‡Department of Medicine, Center for Ulcer Research and Education: Digestive Diseases Research Center, Veterans Administration Wadsworth Center, School of Medicine, University of California Los Angeles, Los Angeles, California
Background & Aims: Enteroglucagon peptides have long been proposed as mediators of intestinal adaptation, including mucosal growth and nutrient absorptive capacity. The hypothesis that infusions of oxyntomodulin, a bioactive form of enteroglucagon, would stimulate glucose and amino acid uptake was tested and its effects were compared with those of glucagon. Methods: Rats were infused intravenously via minipumps with either saline, rat oxyntomodulin (0.47 nmolrkg01rh01), or glucagon (0.88 nmolrkg01rh01) for 7 days, and plasma hormone levels were measured. At death, intestinal dimensions and brush border uptake of D-glucose and L-proline were measured using an in vitro everted sleeve technique. Results: Plasma enteroglucagon and glucagon levels were increased 4and 12-fold, respectively, but there were no effects on food intake, body weight, or intestinal dimensions. In contrast, oxyntomodulin and glucagon significantly stimulated total intestinal glucose uptake capacity by 44% and 53%, respectively, over controls. Oxyntomodulin most potently enhanced glucose uptake in the ileum (215%), whereas glucagon’s greatest effect was in the jejunum (63%–85%). However, neither peptide affected proline uptake. Conclusions: These results support a new, specific action for oxyntomodulin in intestinal adaptation as a glucose uptake stimulator and confirm glucagon’s role as a regulator of glucose uptake.
O
ne of the remarkable properties of the small intestine is its ability to adjust or ‘‘adapt’’ absorptive function under varying dietary, developmental, environmental, or pathological conditions.1 For glucose and amino acids, numerous studies indicate that the specific brush border transporters for those nutrients appear to be regulated by dietary levels of their transported substrate.2 The correlation between dietary nutrients and nutrient uptake rate at once leads to the reasonable suggestion that luminal nutrients might act as the proximate signal regulating transporter activity in the enterocyte. In fact, there is little or no direct evidence for a ‘‘lacoperon’’ model3 in which nutrients themselves, taken up from the gut lumen or blood, modulate transporter expression at the gene or protein levels. Ferraris and / 5e1d$$0005
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Diamond4 clearly pointed out the striking differences in transporter and enzyme regulation between bacterial and intestinal cells. Instead, nutrient transport in enterocytes likely responds to an as yet poorly defined combination of substrate, neuronal, and endocrine signals. Our focus is on the hormonal role of the regulatory gut peptide oxyntomodulin (OXN) in the adaptive regulation of brush border glucose uptake. OXN is one of four major peptides (glicentin and glucagon-like peptide 1 and 2 are the others) derived posttranslationally from a single proglucagon precursor5–7 and cosecreted by L cells in the distal small intestine and colon.8,9 We chose to study OXN as a possible mediator of intestinal adaptation for three reasons. First, numerous studies link elevations in OXN and glicentin levels (known collectively as enteroglucagon, because both enteric peptides contain the entire sequence of glucagon) to circumstances characterized by enhanced nutrient absorption, such as diabetes,10 lactation,11 cold adaptation,11 and recovery from intestinal resection.11 – 13 Because the gut hypertrophies in these conditions, enteroglucagon has long been considered a potential mucosal growth factor.14 In addition, a well-documented up-regulation of glucose transporters per enterocyte occurs along with mucosal growth in some cases (e.g., diabetes).15 Thus, OXN also might increase the activity of specific nutrient transporters. Second, OXN is released in response to a complex set of signals, including luminal nutrients (glucose and fatty acids),16 – 18 neurotransmitters,19 and peptides.19 – 21 Hence, OXN is in the likely chain of signals leading from dietary nutrients to adaptation. Third, no studies have tested infusions of synthetic rat OXN on nutrient uptake. Substantial evidence exists for only one physiological action of OXN, the inhibition of acid secretion (oxyntic cell modulator), the effect for which OXN was named.22 – 24 The three rationales in the preceding paragraph might equally apply to all four major proglucagon peptides; Abbreviations used in this paper: carb-free, carbohydrate-free; GLI, total glucagon-like immunoreactivity; OXN, oxyntomodulin; SGLT1, Na//glucose cotransporter 1. q 1997 by the American Gastroenterological Association 0016-5085/97/$3.00
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however, we focus on OXN in this study for an additional, practical reason. Our recent chemical characterization and synthesis of rat OXN allowed us to perform in vivo infusion tests.25 Rat OXN differs in sequence from commercially available porcine OXN at position 33 (rat, arginine; pig, lysine)7,25 but is identical to presumptive human OXN predicted from the human preproglucagon gene sequence.26 Thus, our first goal was to infuse rats with a synthetic duplicate of their endogenous OXN peptide and measure changes in mucosal growth and brush border D-glucose uptake (mediated by the Na//glucose cotransporter 1 [SGLT1])27 along the intestine. Our infusions targeted OXN elevations at physiological levels observed in animals undergoing intestinal adaptation.14 We also measured uptake of L-proline, mediated by a different Na/dependent (imino) transporter, to test the nutrient specificity of adaptation. Our second goal was to compare effects of OXN infusions with those of glucagon for two reasons: except for an octapeptide extension at its carboxyl terminus, OXN is identical to glucagon in structure, and older studies28–30 have long suggested that glucagon stimulates nutrient absorption, along with recent experiments indicating that glucagon mediates short-term (minutes to days) upregulation of glucose uptake.31,32 This reasoning would similarly suggest that we examine the effects of glicentin infusions, because it too contains the entire sequence of glucagon. However, glicentin’s size (69 amino acids) makes it impractical to synthesize and it is not commercially available. Hence, our studies were limited to OXN and glucagon. We tested both peptides in rats fed a carbohydrate-free (carb-free) diet to assess whether glucose uptake could be stimulated by hormones in the absence of dietary levels of the transported substrate. The results provide the first evidence for a role of OXN in intestinal adaptation, one distinct from enteroglucagon’s role in intestinal growth postulated 25 years ago,33 and confirm and extend glucagon’s action at physiological plasma levels as a regulator of glucose uptake.
Materials and Methods Materials Chemicals (reagent grade) were purchased from commercial sources except where noted. Porcine pancreatic glucagon (ú95% purity) used for infusions and synthetic rat (human, bovine, and porcine) glucagon for radioimmunoassay (RIA) were from Sigma Chemical Co. (St. Louis, MO). Rat OXN was synthesized in the Peptide Biochemistry Core Facility of the CURE: Digestive Diseases Research Center (Los Angeles) as described previously.25 All radioisotopes were from Dupont NEN (Wilmington, DE).
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Animals Male Sprague–Dawley rats (200–250 g) obtained from Sasco, Inc. (Omaha, NE) initially were fed rodent chow (Teklad Rodent Diet; Harlan Teklad, Madison, WI) for 1 week after arrival and then switched to Custom Karasov Carbohydrate-free Diet (ICN Biomedicals, Aurora, OH) 2 weeks before surgery and throughout infusion treatments (described in the following paragraph). Laboratory chows typically are high in carbohydrate (approximately 48%),34 which elevates intestinal glucose uptake rates.4,35 The carb-free diet served to depress basal rates of glucose transport (see Results). All animals were housed singly under a 12/12-hour light-dark cycle at 227C. They received food and water ad libitum throughout the experiment and were used under a protocol approved by the Texas Tech University Animal Care and Use Committee.
Peptide Infusions A total of 24 rats were divided into two sets of 12 rats (6 matched pairs for OXN and control infusions and 6 pairs for glucagon and its control infusions). The infusion pairs were matched closely for body weight (an important determinant of intestinal mass), bled at same time at each sampling period (see next paragraph), and killed together to minimize potential circadian variations in endogenous peptide levels and in nutrient transport rates. For 7 days, 1 rat in each pair received intravenous infusions of 0.9% saline (controls) and the treated rat received either OXN or glucagon (0.47 or 0.88 nmolrkg01rh01, respectively, dissolved in saline) via osmotic minipumps (1 mL/h, Alzet Model 2001; Alza Science Products, Palo Alto, CA). From preliminary experiments, we chose the OXN dosage to increase plasma values of enteroglucagon to about 200 pmol/L. This value represented physiological levels measured in cold-acclimated rats,11 a treatment known to increase glucose absorption.36 The twofold-higher glucagon dosage was selected to achieve an equimolar increase in plasma glucagon, given that the metabolic clearance rate of glucagon is twice that of OXN in sham-operated rats.37 Two of the 6 glucagon-control pairs were removed from the study because of blocked blood-sampling catheters, yielding a final sample size for all measurements of 4 glucagon-control pairs and 6 OXN-control pairs.
Surgical Procedures and Blood Sampling Rats were anesthetized intramuscularly with a solution (1 mL/kg body wt) of ketamine HCl (100 mg/mL; ParkeDavis, Morris Plains, NJ) and xylazine (20 mg/mL; Miles Laboratories, Shawnee, KA) mixed in a ratio of 87:13, respectively. A sterile minipump prefilled with either 0.9% saline or peptide was implanted subcutaneously (SC) 2.5 cm to the right of the sternum and its attached Silastic catheter (0.025 ID 1 0.037 OD; Dow Corning, Midland, MI) tunneled SC to the left external jugular, where it was inserted and secured with 5-0 silk suture for venous infusion. A blood-sampling catheter was then implanted as described by Beverly et al.38 to monitor hormone levels. Briefly, the right exterior jugular vein was exposed via a midline incision in the neck and a 4-cm Silastic
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catheter segment inserted toward the heart. The catheter was secured, tunneled SC, and exteriorized at the top of the head through an L-shaped piece of 21-gauge steel tubing, fixed in position with dental acrylic cement and stainless steel screws anchored in the skull. Catheters were maintained patent by filling with a 40% polyvinylpyrrolidone solution containing 50 U heparin/mL and capped with a heat-sealed piece of Tygon tubing. Blood was sampled from unrestrained rats by first removing the polyvinylpyrrolidone-heparin solution from the catheters and then attaching a 15–20-cm length of Tygon tubing, filled with sterile, heparinized saline to the stainless steel cannula. Blood samples (1 mL) were withdrawn from the rat and mixed immediately with a 100-mL solution containing the anti-peptidase aprotinin (500 U Trasylol/mL blood; FBA Pharmaceuticals, New York, NY) and 1.2 mg Na2 ethylenediaminetetraacetic acid. Plasma was separated by centrifugation (3000g at 47C) and stored at 0807C. Blood was collected during surgery (just before minipump implantation); on infusion day 2, 4, and 6 after surgery; and at death (day 7), with all samples taken between 1300 and 1800 before lights out to minimize diurnal variations and feeding effects on endogenous hormone levels.
Whole-Animal Measurements and Intestinal Dimensions Food intake was recorded daily and body mass on alternate days throughout the experiment. On infusion day 7, nonfasted rats were anesthetized as described previously and a final blood sample was taken. The peritoneal cavity was opened and the entire small intestine rinsed with cold (approximately 47C) mammalian Ringer’s without glucose (in mmol/L: NaCl, 128; KCl, 4.7; CaCl2 , 2.5; KH2PO4 , 1.2; MgSO4 , 1.2; and NaHCO3 , 20; pH 7.4 at 377C, 290 mOsm). The intestine was excised quickly, stripped of mesentery and fat, flushed again with Ringer’s, and divided into three equal proximal, middle and distal parts. Each part was suspended against a vertical scale with a 5-g weight attached for length measurements. The segments were then everted over a glass rod, and the wet mass of each segment was measured. Four everted ‘‘sleeves’’ 1.5 cm in length were cut off from each end of the three intestinal regions for glucose and proline uptake measurements (two replicate sleeves for each nutrient). Tissues handled during the previous procedures were maintained at 47C by rinsing or immersing in cold Ringer’s. We also estimated the nominal surface area per centimeter, the proportion of intestinal thickness represented by mucosa (% scrapable mucosa), and dry mass per centimeter in each gut region using techniques described by Diamond and Karasov.35
Nutrient Uptake Measurements We measured active D-glucose and total L-proline uptake across the intestinal brush border membrane (not transepithelial fluxes) using the everted intestinal sleeve method of Karasov and Diamond.39 Briefly, everted tissue sleeves were secured by ligatures onto glass rods (diameter, 4–6 mm based on intestinal region), preincubated for 5 minutes in Ringer’s
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gassed with 95% O2 –5% CO2 at 377C, and then incubated by suspending the rod over a stirring bar rotating at 1200 rpm in a test tube filled with Ringer’s. The incubation Ringer’s solution differed from the preincubation solution only in that the former contains either 50-mmol/L unlabeled D-glucose or L-proline and trace amounts of D-[14C(U)]glucose or L-[a14 C(U)]proline as nutrient probes. The 50-mmol/L concentrations were selected because they greatly exceed transporter Michaelis constants (Km ), resulting in maximal uptake rates (Vmax ).40,41 We added L-[1-3H(N)]glucose in glucose incubations to correct simultaneously for D-glucose in the adherent fluid and D-glucose taken up passively. After a 2-minute incubation (when uptake is linear with time),40 glucose uptake tissues were rinsed in ice-cold Ringer’s without glucose for 20 seconds to reduce the adherent fluid correction. For proline incubations, we added [1,2-3H]polyethylene glycol (mol wt, 4000) to correct for proline in the adherent fluid, yielding total (i.e., active plus passive) proline uptake. In this case, the rinse at the end of the 2-minute incubation was omitted because it would introduce error into the adherent fluid correction (L-glucose and D-glucose have the same diffusion coefficient but polyethylene glycol and L-proline do not). Instead, only the tip of the rod was blotted to remove excess fluid after proline incubations. For the uptake measurements of both nutrients, a 1-cm portion of the incubated tissue was then cut off and placed into a liquid scintillation vial, and its wet mass determined. The tissue was then solubilized in 1 mL of Soluene 350 (Packard, Meriden, CT) at 407C overnight. The following day, scintillation cocktail (10 mL, Hionic-Fluor; Packard) was added and radioactivity counted on a dual-label scintillation counter (LS6500; Beckman, Fullerton, CA). Activity in the samples was corrected for variable quenching, converted to disintegrations per minute, and the uptake rates calculated as described by Karasov and Diamond.39 In addition to expressing uptake rates normalized to wet mass in different gut regions, we calculated the total intestinal uptake capacity for each nutrient by determining the product of the mean uptake rate per milligram for each region and the region’s wet mass and then summing these products for all small intestinal regions.
Plasma Hormone Measurements We measured plasma glucagon and enteroglucagon (i.e., glicentin plus OXN) to document the actual concentrations of hormones achieved by the minipump infusions. All plasma samples were first extracted with 4% trifluoroacetic acid and then centrifuged (5000g) for 20 minutes at 47C. The supernatant was passed through a C-18 Sep-Pak cartridge (Waters, Milford, MA) and eluted with 90% acetonitrile in 0.1% trifluoroacetic acid in a final volume of 1 mL. Extracts were dried in a Speed-Vac (Savant, Holbrook, NY), reconstituted in assay buffer (0.05 mol/L phosphate-buffered saline with 2% Protenate [Baxter, Glendale, CA] at pH 7.4), and assayed using the appropriate antibodies. Glucagon extraction recoveries averaged 92%, and plasma values were adjusted accordingly. Plasma glucagon was measured by an RIA em-
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ploying a glucagon-specific antibody directed toward the C terminus of the peptide (antibody O4A, purchased from Dr. R. H. Unger, University of Texas Southwestern Medical Center, Dallas, TX).42 Plasma enteroglucagon levels were measured by a subtraction RIA.43 Briefly, pancreatic glucagon levels are subtracted from the total glucagon-like immunoreactivity (GLI), measured with antibody 4304 (kindly provided by Dr. Jens Holst, University of Copenhagen, Copenhagen, Denmark) that recognizes the three GLI peptides containing the glucagon residues 6–15 (i.e., glucagon, glicentin, and OXN).44 Although the subtraction assay does not distinguish between the large (glicentin) and small (OXN) forms of enteroglucagon, preliminary experiments in OXN-infused rats showed that the enteroglucagon activity was related directly to the OXN infusion dosage (data not shown). In both assays, glucagon serves as standard and label (receptor-grade 125I-glucagon). After a 4-day incubation at 47C, dextran-coated charcoal was used to separate bound from free label before counting both fractions in a GammaTrac 1193 (TM Analytic, Elk Grove Village, IL). Enteroglucagon levels are reported as picomolar glucagon equivalents. Intra-assay and interassay coefficients of variation were as follows: glucagon, 11% { 3% and 14% { 5%; and GLI, 14% { 2% and 16% { 4%.
Statistical Analysis Significant differences in mean values between control and peptide-infused rats were determined using Student’s t test for paired observations. Values of P ° 0.05 for group differences were considered statistically significant.
Figure 1. Plasma hormone levels (means { SEM) in paired rats infused for 7 days with saline or peptides. (A ) Enteroglucagon (glicentin / OXN, expressed as picomolar glucagon equivalents; solid lines ) and glucagon (dashed lines ) levels in OXN-infused (●) or saline-infused (m) rats. *P õ 0.005; n Å 6 OXN-control pairs. (B ) Glucagon levels in glucagon-infused (●) or saline-infused (m) rats. *P õ 0.005; n Å 4 glucagon-control pairs.
Results Plasma Hormone Levels in Infused Rats Plasma hormone levels were measured throughout the infusion period to document peptide concentrations achieved in control (saline-infused) and peptide-infused rats. As seen in Figure 1A, a 7-day infusion of rat OXN (0.47 nmolrkg01rh01) significantly elevated plasma enteroglucagon levels fourfold to a plateau of 149 pmol/L (mean value of days 2–6; P õ 0.005) compared with controls (37 pmol/L, mean for all 7 days). In contrast, plasma glucagon concentrations were unaffected by OXN infusion (see Figure 1A). Rats infused with glucagon (0.88 nmolrkg01rh01) showed a 12-fold elevation in plasma glucagon levels (165 pmol/L, mean of days 2–4; P õ 0.005) compared with essentially flat basal values of 14 pmol/L averaged over the 7-day infusion period (Figure 1B). Enteroglucagon levels were not measured in glucagon-infused rats because of inadequate plasma sampling. Note that the infusion rates chosen for OXN and glucagon resulted in similar molar elevations of the respective infused peptides. For both peptides, the decrease in plasma concentrations on day 7 represents exhaustion of the pump reservoir. / 5e1d$$0005
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Food Intake, Body Weight, and Intestinal Dimensions Neither OXN nor glucagon elicited significant differences in food intake compared with controls. Food intake fell to a minimum 24 hours after pump implantation in all groups and then rapidly recovered such that feeding rates on infusion day 7 were equivalent to presurgical values (Figure 2A and B). Consistent with the food intake data, there were no significant differences in body weight between control and peptide-infused rats (Figure 2A and B). We measured intestinal dimensions using multiple types of whole-intestinal and regional tissue techniques to test for effects on intestinal growth. Neither peptide significantly affected whole-intestinal dimensions (Figure 3A and B) nor regional gut mass per centimeter (Figure 4A and B). Other measures of regional tissue dimensions (nominal surface area per centimeter, percent scrapable mucosa, and dry mass per centimeter) similarly showed no changes in response to peptide treatments (data not shown). WBS-Gastro
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Effects on Nutrient Uptake Differences in nutrient uptake per centimeter intestine might result from nonspecific alterations in the number of enterocytes per centimeter (reflected in the milligram wet mass per centimeter value) or from cellspecific changes in transport rate per enterocyte (reflected in the uptake rate per milligram of tissue). Because there were no differences in wet mass per centimeter along the gut (Figure 4A and B), uptake rates were normalized per milligram wet mass. As an additional check for the nutrient specificity of the response, we measured the uptake of two nutrients D-glucose and L-proline, both transported across the brush border primarily by Na/-dependent nutrient transporters.4 Finally, we describe the total uptake capacity for each nutrient to assess the integrated contributions of different gut regions to the whole-intestinal response to OXN and glucagon.
sively in a proximal-distal pattern (Figure 5A). Compared with control values, uptake per milligram was stimulated 12% in the duodenum (P ú 0.05), 30%– 40% in the jejunum (P ú 0.05), and 215% in the ileum (P õ 0.001). Glucagon-infused rats also showed elevated glucose uptake per milligram, but the regional response was distinct from that of OXN-infused rats (Figure 5B). Uptake was increased strongly over control values by 67% (P õ 0.01) in the duodenum, 63%–85% in the jejunum (P õ 0.01), but only 41% in the ileum (P ú 0.05). In contrast to glucose transport, there was no effect of either peptide on proline uptake per milligram (Figure 6A and B). Small Intestinal Nutrient Uptake Capacities
The fourfold elevation in plasma OXN levels achieved after OXN infusions stimulated active glucose uptake, with the strength of response increasing progres-
Figure 7 shows the effects of peptide infusions on the total intestinal uptake capacity for glucose and proline. Saline-control rats on the carb-free diet had glucose uptake capacities of approximately 16 mmol/day. OXN and glucagon infusions significantly increased the uptake capacity for glucose by similar increments over controls (44% and 53%, respectively) to levels of 24 mmolr
Figure 2. Body weight (BWT) and food intake (FI) in (A ) OXN- or (B ) glucagon-infused rats compared with paired saline-infused controls, described in Figure 1. There were no significant effects of either peptide on food intake or body weight compared with saline controls. (A and B ) BWT: n, saline-infused; m, peptide-infused. FI: s, saline; ●, peptide. Mean values { SEM.
Figure 3. Dimensions of the small intestine (pylorus to ileocecal junctions) in 7-day infused rats at death (Figure 1). There were no significant effects of either (A ) OXN or (B ) glucagon on gut length or wet mass. (A and B ) h, saline-infused; , peptide-infused. Mean values { SEM.
Gut Regional Response to Peptide Infusions
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day01rrat01 (Figure 7A). For comparison, consider that rats adapted to a chow (high carbohydrate) diet for ú2 weeks show glucose uptake capacities of approximately 30 mmol/day (unpublished observations, Collie, March 1996). Hence, OXN and glucagon infusions provide carb-free–adapted animals with about 80% of the glucose uptake capacity of chow rats. Without peptide infusions, however, carb-free rats have only 53% of the glucose uptake capacity of chow rats. The increase in uptake capacity of peptide-infused rats appears specific for glucose because proline uptake capacity was unaffected by the peptide infusions (Figure 7B).
Discussion When we elevated plasma concentrations of OXN or glucagon to levels previously observed in other gut adaptation models, glucose uptake across the brush border was significantly stimulated in the absence of dietary carbohydrate. The effect was specific for glucose because (1) proline uptake was unaffected and (2) no changes in intestinal dimensions were detected, which would have led to increased transport of both nutrients in parallel.
Figure 4. Small intestinal wet mass of 1-cm everted sleeves used in the nutrient uptake measurements from duodenum (Duod), jejunum (Jejun), and ileum in infused rats. There were no regional effects of OXN (A, ) or glucagon (B, ) on wet mass per centimeter compared with saline control values (h). For each rat, the value in a given region is the average of four sleeves (two each from the proximal and distal ends of each region), with the means of those averaged values ({ SEM) shown for the 6 OXN-control pairs and 4 glucagon-control pairs.
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The gut regional response to the two peptides was different, with OXN promoting uptake most strongly in the ileum and glucagon, in the proximal two thirds of the intestine. Our results suggest that OXN plays a role in intestinal adaptation not as a growth factor but as a stimulator of active glucose uptake. Enteroglucagon and Glucagon Plasma Levels and Responses Our infusions increased OXN and glucagon to approximately equal plasma levels (149 and 165 pmol/ L, respectively). Plasma enteroglucagon levels reported for different intestinal adaptation models bracket our OXN-infused rat values (in pmol/L): fasted, 3345; fastedrefed, 6845; cold-acclimated/hyperphagic, 20011; and, massive small-bowel resected, 500.12 Similarly, our glucagon-infused rat values are found within the physiological range of plasma glucagon values measured in rats
Figure 5. Active D-glucose uptake across the brush border (normalized per milligram wet mass) in different small intestinal regions of 7-day–infused rats. Uptake was measured in everted sleeves for 2 minutes at 50 mmol/L luminal D-glucose, when influx was linear with time. Values are corrected for both passive uptake and adherent fluid. , Peptide-infused rats; h, saline-infused rats. (A ) OXN infusion increased D-glucose uptake in all regions, but the values were statistically significant only in the ileum (**P õ 0.001; n Å 6 OXN-control pairs). (B ) Glucagon infusion also stimulated D-glucose uptake significantly in all regions (*P õ 0.01, n Å 4 glucagon-control pairs), except in the ileum. Mean value { SEM, with the regional value determined from two adjacent sleeves in each region per rat.
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under various conditions (in pmol/L): resting, 29–5746,47; 6-hour cold exposure, 6648; insulin treatment, 11046; intense exercise, 17946; and fasted neonatal, 146–301.49 Thus, the increase in glucose uptake observed in this study was obtained at physiological OXN and glucagon levels. Debnam and Sharp31 showed recently that chronic glucagon injections (50 mg given intraperitoneally three times daily for 3 days) stimulated galactose uptake in isolated, upper, and middle villus jejunal cells by 31% and 74%, respectively, through a phlorizin-sensitive (i.e., SGLT1mediated) pathway. Because glucose and galactose are both transported actively by SGLT1, their data agree closely with our 63%–85% stimulation by glucagon of glucose uptake in intact jejunal tissue. However, it should be pointed out that their dose, used in several earlier studies,28,29,50,51 results in supraphysiological glucagon elevations (approximately 1800 pg/mL or 516 pmol/L).50 OXN and Glucagon Interactions It is clear that the effects of OXN on glucose uptake were not mediated by glucagon, because glucagon
Figure 6. Total L-proline uptake across the brush border (normalized per milligram wet mass) in different small intestinal regions of 7-day– infused rats. Uptake was measured as described for D-glucose (Figure 5) except that uptake is uncorrected for passive transport (see Materials and Methods for details). Unlike D-glucose, L-proline uptake was unaffected by either OXN infusion (A, ) or glucagon infusion (B, ) compared with control values (h). Values and group size are as described in Figure 5.
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levels were not increased in OXN-infused rats (Figure 1A). However, because we did not measure enteroglucagon levels in glucagon-infused rats, the possibility exists that glucagon’s effects resulted from OXN release in response to glucagon infusion. Four observations argue against this possibility. First, the gut regional responsiveness is different for the two peptides (Figure 5), with OXN most effective in the ileum and glucagon in the duodenum and jejunum. Second, previous studies show no effect of glucagon on enteroglucagon or OXN release by the intestine.20,37 Hence, it is unlikely that OXN levels were elevated in our glucagon-infused rats. Third, in most cases the conditions and signals for glucagon and OXN release are quite different. For example, glucagon is released during starvation, exercise, and hypoglycemia,52 whereas OXN is released after a meal and in hyperphagic conditions. The two exceptions are in diabetes 30,52,53 and cold acclimation,48 when both peptides are elevated. In the latter conditions, fuels must be mobilized not only to meet pathological (diabetic glucopenia) or thermoregulatory (cold acclimation) metabolic requirements, but also replenished through increased food intake and absorption. Fourth, glucagon appears capable of directly stimulating glucose uptake in isolated enterocytes.31,32 The rapid time course of this response (within 15 min-
Figure 7. Total intestinal uptake capacity for D-glucose and L-proline in peptide-infused rats. Uptake capacity was calculated by summing the uptake for each region (the mean uptake per milligram multiplied by the milligram of tissue per region) along the small intestine’s length. Both OXN (A, ) and glucagon (B, ) significantly stimulated D-glucose uptake capacity. Note that L-proline uptake capacity was not affected by either peptide. *P õ 0.05; **P õ 0.01.
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utes) supports a direct action of glucagon on enterocytes (discussed in the next paragraph). Thus, OXN and glucagon may play some independent and some complementary roles in enhancing glucose uptake. What might be the physiological significance of OXN’s actions in the ileum, an intestinal region that normally has low rates of glucose absorption?4 Under normal conditions, glucose uptake in proximal gut regions is sufficient to handle ingested glucose loads and OXN levels are not chronically elevated to high levels. However, in circumstances where uptake capacity in the proximal intestine is exceeded (e.g., in hyperphagic conditions, diabetes, intestinal resection, and lactation), glucose uptake in the ileum may become important (additional absorptive capacity is required). It is under these same conditions that OXN levels are elevated, acting as a possible signal for transport up-regulation. By continuous peptide infusions, we tried to reproduce the chronic elevations in OXN associated with such conditions. The resulting increased glucose uptake suggests that OXN is capable of increasing glucose uptake capacity, primarily by stimulating uptake in the distal gut. Mechanisms of OXN and Glucagon Actions on Glucose Uptake The everted sleeve technique used in this study measures active glucose uptake39 across the brush border membrane, a process mediated by the transporter SGLT1.54 How might OXN or glucagon stimulate SGLT1 transport of glucose? Both indirect and direct mechanisms are feasible. One or both peptides might act indirectly by releasing other peptides or chemical mediators that directly stimulate glucose uptake. For example, both OXN and glucagon stimulate release of insulin,55 which has been reported recently to exert short-term up-regulation of SGLT1-mediated uptake.56 However, OXN in particular is a weak insulin releaser55 and is considered physiologically insignificant compared with other intestinal incretins7 (e.g., glucagon-like peptide 1 and gastric inhibitory peptide). Thus, although insulin cannot be ruled out as a mediator for glucagon-induced uptake stimulation, it is unlikely to mediate OXN’s effects at the dosages used in this study. A second example of indirect signals might involve hyperglycemia, because glucagon and OXN (at sufficiently high doses) elevate blood glucose levels.55 Hyperglycemia per se is thought to signal increased glucose absorption by an unknown mechanism.2,41 Although we did not measure blood glucose levels in this study, infusions of OXN or glucagon at higher dosages (up to 3 nmolrkg01rh01) did not chronically result in blood glu/ 5e1d$$0005
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cose values above 100 mg/dL (unpublished observations, Collie and Mahaney, 1996). Thus, hyperglycemia probably did not signal the enhanced glucose uptake at the lower dosages described in this report. Direct mechanisms of action on enterocytes themselves would potentially include adenosine 3*,5*-cyclic monophosphate (cAMP) elevations mediated through separate OXN and glucagon receptors. Isolated rat jejunal enterocytes incubated with glucagon (10 nmol/L) show increased glucose uptake within 15 minutes.31 Glucagon incubation (0.1–100 nmol/L) caused dose-dependent increases in intracellular cAMP concentration of these cells over a similar time course.32 In the latter study, raising cAMP levels by adding dibutyryl cAMP and theophylline similarly stimulated glucose uptake. These findings suggest a direct action of glucagon on enterocytes, one mediated by increased cAMP. Although receptor binding by OXN also leads to cAMP elevations, the OXN receptor is probably distinct from that for glucagon, as indicated by differing receptor affinities for the two peptides in different target tissues. OXN is 20-fold more potent than glucagon in elevating cAMP levels in isolated rat gastric glands57 but binds to hepatic glucagon receptors with a 50-fold lower affinity than glucagon.55 Our finding that OXN has a regional profile of glucose uptake stimulation distinct from glucagon’s is consistent with the hypothesis of separate receptors. Along with the recent cloning of the glucagon receptor, Jelinek et al.58 showed that glucagon binding causes intracellular elevations not only in cAMP but also in Ca2/. Both second messengers might influence glucose uptake in multiple ways, by stimulating SGLT1 transcription, stabilizing its transcript, or modulating its posttranslational activity. In this regard, the observations that SGLT1 messenger RNA expression is up-regulated by cAMP59 but negatively regulated by protein kinase C activation60 in LLC-PK1 (porcine kidney) cells are particularly interesting. Recent work in diabetic rats, an animal model with elevated enteroglucagon and glucagon plasma levels, indicates that several transporters involved in intestinal sugar absorption (SGLT1 and facilitative glucose transporter 1 and 5) show increased expression at the messenger RNA and protein levels.15,61,62 One additional mechanism proposed for glucagon’s stimulation of Na/-dependent glucose and amino acid uptake is an increased electrochemical gradient for Na/.32,51 However, our data do not support this hypothesis. An Na/ gradient enhanced by either glucagon or OXN would lead to increased uptake of both glucose and proline; yet we observed an increase in glucose uptake alone. Thus, our findings are consistent with the specific induction of SGLT1 by OXN and glucagon. WBS-Gastro
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Future Work Currently, we are testing a wide range of OXN and glucagon doses to establish a full dose-response curve for their effects on glucose uptake. Although we compared the effects of the two peptides in this report at approximately the same plasma levels, the glucagon elevation represents a much larger relative increase (12-fold) than that for OXN (4-fold) over basal values. It is possible that at different concentrations the gut regional pattern of response may be altered or mucosal growth stimulated. However, at the OXN dosages infused, we found no evidence for the mucosal growth effect predicted from previous correlations of gut hypertrophy and enteroglucagon plasma levels. Recently, Gregor et al.63 summarized findings that suggest enteroglucagon may even inhibit enterocyte proliferation. Thus, the connection between this peptide family and mucosal growth regulation remains elusive. Finally, the mechanism of OXN and glucagon action on glucose transport, whether direct or indirect, is currently unclear. The availability of molecular probes for transporter expression and of better cell culture systems, combined with in vivo studies, will help fill in the missing steps that must ultimately link food intake, signal release, and intestinal adaptation.
References 1. Karasov WH, Diamond JM. Adaptive regulation of sugar and amino acid transport by vertebrate intestine. Am J Physiol 1983; 245:G443–G462. 2. Ferraris RP, Diamond JM. Regulation of intestinal nutrient transport. In: Johnson LR, ed. Physiology of the gastrointestinal tract. New York: Raven, 1994:1821–1844. 3. Jacob F, Monod J. Genetic regulatory mechanisms in the synthesis of protein. J Mol Biol 1961;3:318–356. 4. Ferraris RP, Diamond JM. Specific regulation of intestinal nutrient transporters by their dietary substrates. Annu Rev Physiol 1989; 51:125–141. 5. Heinrich G, Gros P, Habener JF. Glucagon gene sequence. J Biol Chem 1984;259:14082–14087. 6. Mojsov S, Heinrich G, Wilson IB, Ravazzola M, Orci L, Habener JF. Preproglucagon gene expression in pancreas and intestine diversifies at the level of post-translational processing. J Biol Chem 1986;261:11880–11889. 7. Holst JJ, Ørskov C. Glucagon and other proglucagon-derived peptides. In: Walsh JH, Dockray GJ, eds. Gut peptides: biochemistry and physiology. New York: Raven, 1994:305–340. 8. Bloom SR, Polak JM. Enteroglucagon and the gut hormone profile of intestinal adaptation. In: Robinson JWL, Dowling RH, Riecken EO, eds. Mechanisms of intestinal adaptation. 1st ed. Lancaster, PA: MTP, 1982:189–198. 9. Jin SLC, Hynes MA, Lund PK. Ontogeny of glucagon messenger RNA and encoded precursor in the rat intestine. Regul Pept 1990; 29:117–131. 10. Heding LG, Rasmussen SM. Determination of pancreatic and gut glucagon-like immunoreactivity (GLI) in normal and diabetic subjects. Diabetologia 1972;8:408–411. 11. Jacobs LR, Bloom SR, Dowling H. Response of plasma and tissue levels of enteroglucagon immunoreactivity to intestinal resection, lactation and hyperphagia. Life Sci 1981;29:2003–2007.
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12. Gornacz GE, Ghatei MA, Al-Mukhtar MYT, Yeats JC, Adrian TE, Wright NA, Bloom SR. Plasma enteroglucagon and CCK levels and cell proliferation in defunctioned small bowel in the rat. Dig Dis Sci 1984;29:1041–1049. 13. Taylor RG, Beveridge DJ, Fuller PJ. Expression of ileal glucagon and peptide tyrosine-tyrosine genes: response to inhibition of polyamine synthesis in the presence of massive small-bowel resection. Biochem J 1992;286:737–741. 14. Bloom SR, Polak JM. The hormonal pattern of intestinal adaptation. Scand J Gastroenterol 1982;72:93–103. 15. Burant CF, Flink S, Depaoli AM, et al. Small intestine hexose transport in experimental diabetes: increased transporter mRNA and protein expression in enterocytes. J Clin Invest 1994;93: 578–585. 16. Unger RH, Ohneda A, Valberde I, Eisentraut AM, Exton J. Characterization of the response of circulating glucagon-like immunoreactivity to intraduodenal and intravenous administration of glucose. J Clin Invest 1968;47:48–65. 17. Jian R, Besterman HS, Sarson DL, Aymes C, Hostein J, Bloom SR, Ramboud JC. Colonic inhibition of gastric secretion in man. Dig Dis Sci 1981;26:195–201. 18. Miazza BM, Al-Mukhtar MYT, Salmeron M, Ghatei MA, Felce-Dachez M, Filali A, Villet R, Wright NA, Bloom SR. Hyperenteroglucagonaemia and small intestinal mucosal growth after colonic perfusion of glucose in rats. Gut 1985;26:518–524. 19. Buchan AMJ, Barber DL, Gregor M, Soll AH. Morphologic and physiologic studies of canine ileal enteroglucagon-containing cells in short-term culture. Gastroenterology 1987;93:791–800. 20. Brubaker PL. Regulation of intestinal proglucagon-derived peptide secretion by intestinal regulatory peptides. Endocrinology 1991; 128:3175–3182. 21. Roberge JN, Gronau KA, Brubaker PL. Gastrin-releasing peptide is a novel mediator of proximal nutrient-induced proglucagonderived peptide secretion from the distal gut. Endocrinology 1996;137:2383–2388. 22. Dubrasquet JM, Bataille D, Gespach C. Oxyntomodulin (glucagon37 or bioactive enteroglucagon): a potent inhibitor of pentagastrin-stimulated acid secretion in rats. Biosci Rep 1982;2:391– 395. 23. Jarrousse C, Audousset-Puech MP, Dubrasquet M, Niel H, Martinez J, Bataille D. Oxyntomodulin (glucagon 37) and its C-terminal octapeptide inhibit gastric acid secretion. FEBS Lett 1985;188: 81–84. 24. Le Quellec A, Kervran A, Blache P, Ciurana AJ, Bataille D. Oxyntomodulin-like immunoreactivity: diurnal profile of a new potential enterogastrone. J Clin Endocrinol Metab 1992;74:1405–1409. 25. Collie NL, Walsh JH, Wong HC, Shively JE, Davis MT, Lee TD, Reeve JR Jr. Purification and sequence of rat oxyntomodulin. Proc Natl Acad Sci USA 1994;91:9362–9366. 26. Bell GI, Sanchez-Pescador R, Laybourn PJ, Najarian RC. Exon duplication and divergence in the human preproglucagon gene. Nature 1983;304:368–371. 27. Hediger MA, Rhoads DB. Molecular physiology of sodium-glucose cotransporters. Physiol Rev 1994;74:993–1026. 28. Rudo ND, Rosenberg IH. Chronic glucagon administration enhances intestinal transport in the rat. Proc Soc Exp Biol Med 1973;142:521–525. 29. Debnam ES. Effect of sodium concentration and plasma sugar concentration on hexose absorption by the rat jejunum in vivo. Pflu¨gers Arch 1982;393:104–108. 30. Riecken EO, Gregor M. Glucagon and small-bowel mucosa. Scand J Gastroenterol 1985;20:30–40. 31. Debnam ES, Sharp PA. Acute and chronic effects of pancreatic glucagon on sugar transport across the brush-border and basolateral membranes of rat jejunal enterocytes. Exp Physiol 1993; 78:197–207. 32. Sharp PA, Debnam ES. The role of cyclic AMP in the control of
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Received August 8, 1996. Accepted January 31, 1997. Address requests for reprints to: Nathan L. Collie, Ph.D., Department of Biological Sciences, Box 43131, Texas Tech University, Lubbock, Texas 79409-3131. Fax: (806) 742-2963. Supported by National Institutes of Health grant DK 42973, a TTU Institute for Nutritional Sciences grant, and a Howard Hughes Medical Institute Fellowship (to S.J.) through the Undergraduate Biological Sciences Education Program. Dr. Zhu’s current affiliation is: Division of Medicine, M. D. Anderson Cancer Center, University of Texas, Houston, Texas 77030. The authors thank Jared Diamond and John Walsh for their support and discussions at critical stages in this project. Wade Mahaney, Richard Lombardini, and Carol Zhang provided valuable assistance in manuscript preparation and graphics.
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