1 Sugar sensing by enterocytes combines polarity, membrane bound 2 detectors and sugar metabolism
1,2,3,#,*1,2,3,#1,2,31,2,34 Maude Le Gall, Vanessa Tobin, Emilie Stolarczyk, Véronique Dalet,
1,2,31,2,3 5 Armelle Leturque, Edith Brot-Laroche
7 1 INSERM, UMR S 872, Centre de Recherche des Cordeliers, Paris F-75006 France. 8 2 Université Pierre et Marie Curie-Paris6, UMR S 872, Paris F-75006 France. 9 3 Université Paris Descartes, UMR S 872, Paris F-75006 France.
#11 MLG and VT have equally contributed to the work.
* Correspondence to : Maude Le Gall, UMRS 872 Centre de Recherche des Cordeliers, 15 rue de 13
14 l’Ecole de Médecine, Paris, F-75006 France. Tel : + 33 1 42 34 68 99. Fax: + 33 1 43 25 16 15. Email: 15 email@example.com
17 Running head : Sugar signalling pathways in enterocytes
18 Keywords: glucose signalling, enterocyte, GLUT2, sugar metabolism, sweet taste receptor. 19 Total number of text figures and tables : 6 figures
20 Funded by: ALFEDIAM Merck Lipha, Institut Benjamin Delessert, AIP ATC Nutrition; 21 Grant Number: ASEO22129DSA.
23 Abbreviations : SI: sucrase isomaltase, L-PK: liver-pyruvate kinase, SGLT1: sodium glucose 24 transporter, GPCR: G protein-coupled receptor, T1R: Taste Receptor type 1, 3OMG: 3-O-25 methyl glucose, N-AGA: N-acetylglucosamine, G-6-Pase : glucose-6-phosphatase, EGFP: 26 enhanced green fluorescent protein, ChREBP: carbohydrate response element binding protein, 27 ChoRE: carbohydrate response element, SREBP: sterol response element binding protein, 28 LXR: liver X receptor, PKA: protein kinase A, ADRP, adipose differentiation-related protein. 29
3 Sugar consumption and subsequent sugar metabolism are known to regulate the expression of 4 genes involved in intestinal sugar absorption and delivery. Here we investigate the hypothesis 5 that sugar-sensing detectors in membranes facing the intestinal lumen or the bloodstream can 6 also modulate intestinal sugar absorption. We used wild-type and GLUT2-null mice, to show 7 that dietary sugars stimulate the expression of sucrase-isomaltase (SI) and L-pyruvate kinase 8 (L-PK) by GLUT2-dependent mechanisms, whereas the expression of GLUT5 and SGLT1, 9 did not rely on the presence of GLUT2. By providing sugar metabolites, sugar transporters, 10 including GLUT2, fuelled a sensing pathway. In Caco2/TC7 enterocytes, we could disconnect 11 the sensing triggered by detector from that produced by metabolism, and found that GLUT2 12 generated a metabolism-independent pathway to stimulate the expression of SI and L-PK. In 13 cultured enterocytes, both apical and basolateral fructose could increase the expression of 14 GLUT5, conversely, basolateral sugar administration could stimulate the expression of 15 GLUT2. Finally, we located the sweet-taste receptors T1R3 and T1R2 in plasma membranes, 16 and we measured their cognate Galpha Gustducin mRNA levels. Furthermore, we showed 17 that a T1R3 inhibitor altered the fructose-induced expression of SGLT1, GLUT5 and L-PK. 18 Intestinal gene expression is thus controlled by a combination of at least three sugar-19 signalling pathways triggered by sugar metabolites and membrane sugar receptors that, 20 according to membrane location, determine sugar sensing polarity. This provides a rationale 21 for how intestine adapts sugar delivery to blood and dietary sugar provision.
3 Intestinal sugar delivery depends on the levels of expression of dissacharidases (i.e. sucrase-4 isomaltase, SI) and sugar transporters. Indeed, dietary glucose is transported across the apical 5 membrane of enterocytes by the energy-dependent sodium-glucose co-transporter 1 (SGLT1) 6 and dietary fructose by the facilitative transporter GLUT5 (Levin, 1994). At the basolateral 7 membrane, glucose and fructose exit the intestinal epithelia mainly via GLUT2-dependent 8 facilitated diffusion (Levin, 1994) to reach the bloodstream. In GLUT2-null mice, glucose 9 delivery seems to be mediated via vesicular trafficking (Stumpel et al., 2001) and the 10 mechanism by which fructose reaches the blood stream is probably dependent on the presence 11 of GLUT5 in the basolateral membrane (Blakemore et al., 1995). In the case of a sugar rich 12 meal, GLUT2 can be recruited into the apical membrane where it complements SGLT1 and 13 GLUT5 transport capacities (Kellett and Brot-Laroche, 2005). Furthermore, the small 14 intestine adapts to repeated sugar ingestion by increasing transporter expression over the 15 course of a few days (Burant and Saxena, 1994; Goda, 2000; Inukai et al., 1993). Thus, the 16 regulation of the transcription, expression and location of sugar transporters organizes sugar 17 absorption. In pathological conditions this regulation can amplify sugar delivery creating a 18 vicious circle. Indeed, in rat, streptozotocin-induced diabetes increases SGLT1, GLUT2 and 19 GLUT5 mRNA levels and sugar absorption (Burant et al., 1994; Ferraris et al., 1993), 20 possibly causing exacerbated postprandial blood glucose excursion. Knowledge of the 21 mechanisms of sugar sensing at the luminal and basolateral enterocyte membranes is thus 22 essential for an understanding of the mechanism of induction of sugar-sensitive genes. 23 The regulation by sugars of gene expression is highly conserved through evolution, and is
found in bacteria (Jacob and Monod, 1961), yeast (Johnston, 1999), plants (Villadsen and 24
25 Smith, 2004), and mammals (Girard et al., 1997; Towle, 2005). In pancreatic ß cells, glucose 26 is known to alter the expression of more than 150 genes that can be grouped in functional 27 clusters, including insulin secretion, energy metabolism, membrane transport, signalling 28 pathways, gene transcription and protein synthesis/degradation (Schuit et al., 2002). Fructose 29 is known to regulate 50 genes in the small intestine of neonate rats, where it affects genes 30 encoding for ion and hexose-transporters as well as enzymes of hexose metabolism (Cui et al., 31 2004).
32 Considering the number and functional variety of sugar regulated genes, and given their 33 location in diverse sugar-sensitive tissues, a single signalling pathway is unlikely to explain
1 all cellular adaptations to life in a sugar-containing environment. Our current understanding 2 of how cells respond to sugar is mainly based on studies in yeast and involves signalling 3 pathways either initiated from hexose metabolites directly or from the direct engagement of 4 hexoses with their cognate sugar receptors at the cell surface (Holsbeeks et al., 2004). In 5 mammals, studies have been primarily focused on glucose metabolism and related signals 6 (Girard et al., 1997; Towle, 2005). However, non metabolisable sugar analogues can stimulate 7 SGLT1 expression (Miyamoto et al., 1993). Some sugar receptors have been identified in the 8 plasma membrane of mammalian cells. A first group of mammalian sugar receptors is 9 composed of sugar transporters, which can also function as sugar detectors. This group 10 includes GLUT2 and SGLT3. Indeed, in hepatocytes GLUT2 can fuel sugar metabolism and 11 also trigger a receptor-dependent protein signalling cascade (Guillemain et al., 2000; 12 Guillemain et al., 2002). SGLT3 functions in cholinergic neurons neighboring enterocytes 13 and while it does not transport glucose, it induces membrane currents upon sodium-dependent 14 glucose binding (Diez-Sampedro et al., 2003). A second group of sugar detectors contains 15 members of the G Protein–Coupled Receptor (GPCR) family. Among them are the sweet
16 taste receptors of type 1, T1R, located on the tongue. Heterodimeric T1R3/T1R2 and T1R3 17 homodimers form sweet taste receptors that bind fructose or sucrose, leading to adenylate-18 cyclase signalling cascades (Nelson et al., 2001; Xu et al., 2004; Zhao et al., 2003). 19 Biochemical data suggest that GPCRs participate to sugar signalling in an enteroendocrine 20 cell line, but their molecular identification has yet to be determined (Rozengurt, 2006). In 21 enterocytes, cell polarity further complicates the regulation of sugar sensitive genes. Indeed, 22 detectors may conceivably be activated by dietary sugars at the apical membrane and by 23 blood glucose at the basolateral membrane of enterocytes.
24 In this study we consider both metabolism-driven sugar signalling and sugar detection 25 initiated signalling pathways. By monitoring the induction of sugar sensitive genes both in
26 vivo and in Caco-2/TC7 cells we report here the identification of distinct sugar sensing 27 signalling pathways with a striking polar distribution in enterocytes. These sugar sensory 28 mechanisms offer insight not only into the modes of sugar absorption and delivery, but also 29 unveil important avenues for the development of novel pharmacological compounds for 30 improved glycaemic control.
1 MATERIALS AND METHODS
3 Mice: Wild-type mice were from C57Bl/6 strain (Janvier, France). GLUT2-null mice (RIP 4 GLUT1XGLUT2-/-) (Guillam et al., 1997) were bred in the transgenic animal facility of 5 IFR58 (Paris, France). All animal procedures are complied with recommendations for the use 6 of laboratory animals from the French administration. Mice were fed for 5 days with the 7 experimental diets, which contained either low amounts of sugar, or 65% (W/W) glucose or 8 fructose as previously described (Gouyon et al., 2003a). The jejunum is here defined as the 9 part of the small intestine starting at the Treitz ligament and excluding its last third. Intestinal 10 samples, taken from fed mice, were rapidly everted and contents washed out in ice cold PBS 11 before mucosa scrapings were taken. Mucosa was dispersed in RNA extraction buffer and 12 snap frozen in liquid nitrogen.
5214 Cell culture: Caco-2/TC7 cells were seeded at 6 x10 cells/cm either on six-well, solid or
15 porous (3µm high pore density) supports (Becton Dickinson, Meylan, France). Cells were
-116 grown in complete Dulbecco’s modified Eagle’s medium (25 mmol.L glucose DMEM,
17 Gibco, Paisley, U.K) supplemented with 20% heat-inactivated (30min, 56?C) fetal calf serum 18 (FCS) (AbCys, Paris, France). Media were renewed every 24 hours for at least 10 days to 19 allow differentiation of the cells (Chantret et al., 1994; Mahraoui et al., 1994). 20 Post-confluent, differentiated cells were switched from standard growth media (DMEM 25
-121 mmol.L glucose) to glucose-free DMEM supplemented with 10% heat-inactivated FCS, and
-122 contained less than 1 mmol.L glucose (low sugar medium). According to need, media were
23 supplemented with sugar for 2 to 4 days as indicated in the legend of the figures. For sugar
-1 -1metabolim assay; 25 mmole.L3-O-Methylglucoside or 250 mmole.L24 N-AGA were added
25 at both apical and basal sides of the cells for 48 hours. For sweet taste receptor functional 26 assay, after differentiation, cells were grown in glucose and glutamine free DMEM
-127 supplemented with 10% dialyzed FCS. Inhibitor, 1 mmol.L lactisole (Sigma Aldrich, Saint
-128 Quentin, France) and substrate 25 mmol.L fructose, were added during the last 48 hours of
29 culture at both apical and basal poles.
30 When grown on porous support, cell viability and cell monolayer integrity after treatments 31 were estimated by measure of the transepithelial electrical resistance (TEER), which is a 32 witness of tight junction integrity and of ion pump function in cell membranes (Grasset et al., 33 1984).
1 Immunofluorescence: Differentiated Caco-2/TC7 cells grown on filters were fixed with 4% 2 Paraformaldehyde (Sigma Aldrich, Saint Quentin, France), permeabilized with 0.2% Triton 3 (Sigma Aldrich, Saint Quentin, France) and labeled with antibodies targeting T1R2 (T-20: sc-4 22456) and T1R3 (N-20: sc-22458) (Santa Cruz, Biotechnology, Tebu France). Images were 5 produced by confocal microscopy (Zeiss LSM510 software).
7 Human GLUT2, GLUT5 and sucrase-isomaltase promoter constructs: Caco-2/TC7 cells were
8 transfected with the following promoter regions inserted into p205 plasmid driving the 9 reporter gene luciferase (Rodolosse et al., 1996) : -1100/+300 of the hGLUT2 promoter 10 (generous gift of GI Bell, University of Chicago, Chicago, IL, USA), –3600/+60 of the hSI
11 promoter (Rodolosse et al., 1997) and -2500/+21 of the hGLUT5 promoter (Mahraoui et al., 12 1994). Populations of stably transfected cells were established. Protein assays were made with 13 the BCA kit (Pierce, Interchim, Montluçon, France). Maximal variations of protein 14 concentrations between different cell cultures and different culture conditions were below 15 20%. Luciferase activities were measured using the Luciferase assay kit (Promega, 16 Charbonnières les Bains, France) in a Lumat LB9501 luminometer (Berthold Detection
-1-117 System, Pforzheim, Germany). Results were expressed as relative light units RLU.sec.µg
20 EGFP-GLUT2-loop and -C-terminus peptide constructs: The coding regions of the
21 intracellular loop between transmembrane domain 6 and 7 (amino acids 237-301) and the C-22 terminus GLUT2 domain (amino acids 481-521) of rat GLUT2 were cloned in frame with 23 EGFP in pEGFP-C (Clontech BD Biosciences, le Pont de Claix, France), as previously
described (Guillemain et al., 2000). The EGFP-GLUT2 domain cassettes were placed 24
25 downstream the SV40 promoter of pGL3 (Promega, Charbonnières les Bains, France), 26 allowing the expression of EGFP-GLUT2 domains in differentiated Caco-2/TC7 cells. Stably 27 transfected EGFP positive cells were established and sorted by FACS (Epics Altra, Beckman 28 Coulter, Roissy, France). The cells were secondarily and transiently transfected with the 29 hGLUT2 promoter using the lipofectin transfection kit (Life Technologies, Cergy Pontoise, 30 France).
32 Messenger RNA: Total RNA from the jejunum of mice or from Caco2/TC7 cells were 33 extracted using TriReagent (MRC, Interchim, Montluçon, France). Mouse and human
1 GLUT2, L-Pyruvate kinase (L-PK), Glucose-6-phosphatase (G-6-Pase), Sucrase-Isomaltase 2 (SI) and human SGLT1 and GLUT5 mRNAs were quantified by reverse transcription and
3 real-time PCR using the Light-Cycler System according tothe manufacturer's procedures
4 (Roche Molecular Biochemicals, Meylan, France) as previously described (Gouyon et al., 5 2003a; Guillemain et al., 2002). The primers used were for hGLUT5 forward 5’-
6 TCTCCTTGCAAACGTAGATGG-3’and reverse 5’-GAAGAAGGGCAGCAGAAGG-3’, for hSGLT1
7 forward 5’-TGGCAATCACTGCCCTTTA-3’and reverse 5’-TGCAAGGTGTCCGTGTAAAT-3’ and for
8 hADRP forward 5’-GTGAGATGGCAGAGAACGGTGTG- 3’and reverse 5’-
9 TGCCCCTTTGGTCTTGTCCA-3’. All primer pairs amplified a single amplicon as indicated by the 10 unique melting temperature of the PCR product. Moreover, we verify the size and specificity 11 of the amplicon by restriction enzyme analysis and electrophoresis. The large ribosomal
12 protein L19 was used as a control gene since its expression level varied by less than 20% in 13 all the culture conditions applied to the cells. Results were expressed as ratios of gene mRNA 14 over L19 mRNA levels. Northern blots were performed to measure mouse intestinal GLUT5 15 and SGLT1 mRNA. Density analyses (Gel Analyst 3.02 software) were expressed as the ratio 16 of mRNA to 18S rRNA. For human taste receptors identification, primers were selected using 17 published data (Rozengurt et al., 2006) or predicted sequences available in the NCBI database: 18 NM_152232 for T1R2, NM_152228 for T1R3, and XM_001129050, XM_294370 and
19 X_M939789 for Gustducin. T1R2: forward 5’-GTATGAAGTGAAGGTGATAGGC-3’and reverse
20 5’-GGGTAGACCACCCTCTTGG-3’; T1R3: forward 5’-CAAGTTCTTCAGCTTCTTCCTC-3’ and
21 reverse 5’-GTACATGTTCTCCAGGAGCTGC-3’; Gustducin: forward 5’-
22 GCCAAATACATTTGAAGATGCAGG-3’ and reverse 5’- GCACTTCTGGGATTTACATAATC-3’. Note
23 that for T1R2 and T1R3 nested primers were also used. Nested T1R2: forward 5’-
24 TGCGCTTCGCGGTGGAGG-3’and reverse 5’-CAGCCGAGGAGGCTGTGC-3’; nested T1R3: forward
25 5’-GGTCAGCTACGGTGCTAGC-3’and reverse 5’-AGCCTGAGGCGTTGCACTG-3’.
27 Sugar transepithelial transfer: The glucose or fructose content of the apical and basal culture 28 media was assayed with an enzymatic assay kit (Sigma Aldrich, Saint Quentin, France). The 29 non-specific transpithelial transfer of sugar across Caco-2/TC7 cells grown on porous support
330 was measured using (1-H NEN) L-glucose. L-glucose transfer was lower than 1.5 % of the 31 D-isomers (3 independent experiments, data not shown) indicating that epithelia were tight. 32 Using fructose media helped to distinguish fructose and glucose transporting GLUTs. 33
1 Statistics : All statistical analyses were made using ANOVA and student T tests (PRISM 2 software).
2 Regulation of intestinal genes by dietary sugars
3 In vivo experiments were first conducted to determine if glucose and fructose employ distinct 4 signalling pathways to induce sugar-regulated genes. We focused our study on sugar 5 transporters (GLUT5, GLUT2, SGLT1) and sugar metabolism enzymes (SI and L-PK). As 6 previously observed for GLUT2 (Gouyon et al., 2003a), glucose- or fructose-rich diets 7 efficiently stimulated the mRNA accumulation of these sugar-regulated genes in the jejunal 8 mucosa of wild-type (WT) mice (Figure 1). SGLT1, SI and L-PK mRNA increased to similar 9 levels under both glucose and fructose dietary regimens (Figure 1). On the other hand, 10 GLUT5 mRNA increased in mice fed fructose- but not glucose-rich diets (compare Figure 1A 11 to 1B), in agreement with previous studies (Burant and Saxena, 1994; Gouyon et al., 2003b) 12 indicating that cells discriminate glucose and fructose signals to regulate the expression of 13 GLUT5. As glucose and fructose are both catabolized through glycolysis in enterocytes, 14 metabolic signalling via glycolysis cannot fully account for the differential effect. We used 15 GLUT2-null mice to document the contribution of this glucose/fructose transporter to the 16 regulation of sugar-sensitive genes. The mRNA levels were similar for the different genes 17 analysed in GLUT2-null and WT mice fed low carbohydrate diets. GLUT2-null and WT mice 18 exhibited indistinguishable increases of GLUT5 mRNA in response to fructose-rich diet. 19 Similarly, the small but significant induction of SGLT1 by sugar-rich diets was similar in 20 GLUT2-null and WT mice. By contrast, SI and L-PK mRNA inductions were not observed or 21 were dramatically reduced in GLUT2-null mice. Therefore, a GLUT2-dependent signalling 22 pathway is required to obtain SI and L-PK gene regulation by dietary glucose and fructose. 23 Interestingly, feeding GLUT2-null mice with fructose produced a partial accumulation of L-
PK mRNA (Figure 1B), indicating that fructose and glucose signalling pathways can differ in 24
25 the intestine. These in vivo data indicate that enterocytes exploit several sugar signalling 26 pathways to adapt the expression of sugar target genes to the dietary environment. 27 Effect of sugars on sensitive genes in Caco-2/TC7 enterocytes
28 Enterocyte detection of dietary sugars could occur at their apical membranes exposed to the 29 intestinal lumen content or their basolateral membranes exposed to the blood. To assess the 30 sugar detection capacity of each of these membranes in a controlled manner, we used 31 differentiated human colon carcinoma, Caco-2/TC7 cells that display the morphological 32 characteristics and functional properties of intestinal absorptive cells (Delie and Rubas, 1997).
1 Caco-2/TC7 cells cultured on solid support have only their apical poles in contact with culture 2 media, whereas cultures grown on porous support permit sugar provision to either apical or 3 basolateral membranes. This system was thus employed to differentially expose enterocytes 4 to dietary sugars and to determine polarity in sugar detection and signalling. 5 As shown in Figure 2, fructose stimulated hGLUT5 promoter activity by 2 fold in Caco-6 2/TC7 cells regardless of whether they were grown on solid or porous support. By contrast, 7 neither glucose nor fructose could stimulate the hGLUT2 promoter activity in cells grown on 8 solid support (Figure 2), although the same hGLUT2 promoter fragment has been shown 9 capable of being activated by glucose in pancreatic MIN6 cells (Cassany et al., 2004). 10 However, a net stimulation of hGLUT2 promoter activity was observed in Caco-2/TC7 cells 11 grown on porous support in agreement with the capacity of dietary sugars to induce GLUT2 12 in vivo. Thus growing enterocytes on porous support with their basal poles capable of 13 detecting sugars unmasks polarity in sugar signalling toward GLUT2 gene regulation. 14 We took advantage of the restricted localization of SI at the apical membrane of enterocytes 15 to further study the mechanisms of polarity in sugar detection and signalling. The addition of 16 sucrose into apical medium stimulated hSI promoter activity significantly (Figure 2), but no 17 activation was obtained when sucrose addition was restricted to media exposed to the 18 basolateral membrane. Furthermore, sucrose cannot be simply hydrolyzed to fructose and 19 glucose at the basolateral membrane as it lacks SI activity. These data also indicate that 20 epithelial tight junctions were functional. Importantly, the glucose and fructose moieties of 21 sucrose stimulated the hSI promoter activity irrespective of the compartment of addition 22 (Figure 2). Thus, enterocytes growing on porous support display polarity in their responses to 23 dietary sugars.
24 When administered independently to the apical or basal compartments, glucose and fructose 25 concentrations equilibrated between the apical and basolateral media within 12 hours (Figure 26 3A). Therefore, in our culture conditions, polarized sugar signals could only be detected 27 within 12 hours (i.e. before sugar equilibration is achieved). We measured the time course of 28 hGLUT2 promoter activity induced by sugar (Figure 3B, left panel) and unfortunately, the 29 addition of sugar required 48 hours to maximally stimulate the hGLUT2 promoter and little to 30 no stimulation was observed at 12 hours (Figure 3B, left panel). Accordingly, the addition of 31 glucose to either pole of the cell for 48h (i.e. long after sugar equilibration is achieved), 32 produced a similar stimulation of the hGLUT2 promoter (not shown).