The inhibitory effects of the angiotensin-converting enzyme (ACE)-ANG II-angiotensin type 1 (AT1) receptor axis on jejunal glucose uptake and the reduced expression of this system in type 1 diabetes mellitus (T1DM) have been documented previously. The ACE2-ANG-(1–7)-Mas receptor axis is thought to oppose the actions of the ACE-ANG II-AT1 receptor axis in heart, liver, and kidney. However, the possible involvement of the ACE2-ANG-(1–7)-Mas receptor system on enhanced jejunal glucose transport in T1DM has yet to be determined. Rat everted jejunum and Caco-2 cells were used to determine the effects of ANG-(1–7) on glucose uptake and to study the ACE2-ANG-(1–7)-Mas receptor signaling pathway. Expression of target gene and protein in jejunal enterocytes and human Caco-2 cells were quantified using real-time PCR and Western blotting. T1DM increased jejunal protein and mRNA expression of ACE2 (by 59 and 173%, respectively) and Mas receptor (by 55 and 100%, respectively) in jejunum. One millimolar ANG-(1–7) reduced glucose uptake in jejunum and Caco-2 cells by 30.6 and 30.3%, respectively, effects that were abolished following addition of 1 μM A-779 (a Mas receptor blocker) or 1 μM GF-109203X (protein kinase C inhibitor) to incubation buffer for jejunum or Caco-2 cells, respectively. Finally, intravenous treatment of animals with ANG-(1–7) significantly improved oral glucose tolerance in T1DM but not control animals. In conclusion, enhanced activity of the ACE2-ANG-(1–7)-Mas receptor axis in jejunal enterocytes is likely to moderate the T1DM-induced increase in jejunal glucose uptake resulting from downregulation of the ACE-ANG II-AT1 receptor axis. Therefore, altered activity of both ACE and ACE2 systems during diabetes will determine the overall rate of glucose transport across the jejunal epithelium.
- glucose transport
- angiotensin-converting enzyme 2
- Mas receptor
- sodium-dependent glucose transporter 1
- diabetes mellitus
there is increasing evidence that a local renin-angiotensin system (RAS) exerts paracrine, autocrine, and/or intracrine actions in modulating a wide range of physiological functions (26). We have recently proposed that an enterocyte RAS regulates glucose uptake across the small intestinal epithelium. Thus, the binding of angiotensin II (ANG II) to its type 1 (AT1) receptor at the brush-border membrane (BBM) inhibits dose-dependently sodium-dependent glucose transporter 1 (SGLT1)-mediated glucose transport across this membrane (34). The stimulatory effect of type 1 diabetes mellitus (T1DM) on intestinal glucose transport is well documented (7, 8), and this response is likely to contribute to postprandial hyperglycemia in diabetic patients. T1DM promotes both SGLT1- and glucose transporter 2 (GLUT2)-mediated glucose transport at the BBM. However, the mechanisms responsible for increased enterocyte glucose transport in diabetes are unclear but are likely to involve both systemic and local influences. Our recent studies show that streptozotocin (STZ)-induced T1DM was associated with decreased expression of enterocyte RAS components, including angiotensin-converting enzyme (ACE) and AT1 receptor (33). Therefore, downregulation of the enterocyte RAS in diabetes might explain, at least in part, the enhanced glucose uptake seen in this condition (33).
ACE2 is an exopeptidase that is structurally similar to ACE but has different affinities for the two enzymes, making it resistant to inhibition by ACE inhibitors. ACE2 catalyzes the conversion of ANG II to ANG-(1–7) and also the conversion of ANG I to ANG-(1–9) (13). The conversion of ANG II to ANG-(1–7) is the major action of ACE2, since the binding affinity of ACE2 for ANG II is some 400-fold higher than that for ANG I (27, 32). In rats, ACE2 is expressed in many tissues, including the brain (14), heart (15), liver (24), lung (16), and pancreas (30). In human, ACE2 has also been detected in heart and kidney (12) as well as colon and small intestine (31). ANG-(1–7) is an endogenous ligand for the G protein-coupled Mas receptor. Evidence suggests that the ACE2-ANG-(1–7)-Mas receptor axis opposes the local actions of the ACE-ANG II-AT1 receptor axis in liver, heart, and kidney (2). Indeed, the ACE2-ANG-(1–7)-Mas receptor system may compensate for the cellular effects of altered expression of the ACE-ANG II-AT1 receptor system in diabetes (2), as well as in other pathological conditions such as inflammation and fibrosis (29). More recently, a study of ACE2 gene therapy targeting the islet cells has shown that increased ACE2 activity opposes the effects of raised ACE activity (1).
In light of these findings, we hypothesize that an ACE2-ANG-(1–7)-Mas receptor axis in jejunal enterocytes may compensate for the downregulation of the ACE-ANG II-AT1 receptor axis in diabetes, thus providing a moderating influence on glucose uptake. We tested this idea by examining the enterocyte expression and regulation of the ACE2-ANG-(1–7)-Mas receptor system and its role in the control of glucose uptake during increased extracellular glucose levels using an established rat model of T1DM together with the human Caco-2 cell line.
MATERIALS AND METHODS
The study used male adult Wistar rats (280–320 g) supplied by the Laboratory Animal Services Centre at The Chinese University of Hong Kong. All procedures were approved by the Animal Ethical Committee of the Chinese University of Hong Kong. Experimental T1DM was induced by a single tail-vein injection of STZ (80 mg/kg), and animals were used 2 wk later. Control animals were injected with the vehicle (0.1 M citrate buffer, pH 4.5). Animals used for uptake studies were maintained on a maintenance chow (rolab RMH 2500, 5P14; Lab Diet) and water ad libitum up to the time of experimentation. Only those diabetic rats with blood glucose levels above 18 mM were used for the study (33). Anesthesia before experimentation was achieved with pentobarbitone sodium (50 mg/kg ip).
Isolation of enterocytes.
Enterocytes were prepared from 20-cm-long segments of jejunum beginning 10 cm distal to the ligament of Treitz. Villus cells were harvested by a Ca2+-chelation technique (11) that produces enterocytes with a high viability. Briefly, isolated intestinal segments were flushed through with ice-cold saline followed by air. The segment was tied off at one end and filled with Ca2+-free hypertonic isolation buffer (in mM: 7 K2SO4, 44 K2HPO4, 9 NaHCO3, 10 HEPES, 2 l-glutamine, 0.5 dithiothreitol, 1 Na2EDTA, and 180 glucose, pH 7.4), equilibrated with 95% O2-5% CO2, avoiding overdistention. The segment was then tied off to form a closed sac and incubated in 0.9% saline at 37°C with gentle shaking for 16 min. Cells were dislodged manually, and the resulting suspension was collected and centrifuged for 30 s at 500 g. The pellet was resuspended in freshly prepared cold buffer and recentrifuged, a procedure that was repeated two times (33, 34).
Western blot analysis.
The methods used for immunoblotting have been described previously (17). Protein from the enterocyte suspension was extracted using the CytoBuster Protein Extraction Reagent (Novagen, Darmstadt, Germany) and quantified using a Bradford assay kit (Bio-Rad, Munich, Germany). Proteins (10 μg/lane) were subjected to electrophoresis on a 10% (wt/vol) polyacrylamide gel. Protein was transferred from the polyacrylamide gel to the polyvinylidene difluoride membrane using a Semi-Dry Transblot unit (Bio-Rad). The protein blotted membrane was saturated by submersion in 5% (wt/vol) nonfat skimmed milk in PBS (pH 7.4) with 0.1% (vol/vol) Tween 20 for 1 h at room temperature. The membranes were incubated with anti-Mas receptor rabbit polyclonal antibodies (ab66030; Abcam) (1:200), anti-ACE2 rabbit polyclonal antibodies (SC-20998; Santa Cruz Biotechnology) (1:300), and anti-β-actin mouse polyclonal antibodies (Chemicon) (1:5,000) overnight at 4°C. After being rinsed in PBS, the membranes were incubated with the following corresponding peroxidase-labeled secondary antibodies for 1 h at room temperature: anti-rabbit IgG antibody (Amersham) (1:1,300) and anti-mouse IgG antibody (Amersham) (1:2,500). The positive signal was revealed using ECL-plus Western blotting detection reagent and autoradiography film (Amersham). The intensity of the bands was quantified using FlourChem software.
Real-time PCR analysis.
Quantitative RT-PCR was performed using an ABI PRISM 7700 Sequence Detection System (PE Applied Biosystems, Foster City, CA) as described previously (6). Briefly, total RNA was extracted from freshly prepared enterocytes or Caco-2 cells grown in 5.6, 11.2, or 25 mM glucose using Trizol reagent (GIBCO, Invitrogen) according to the manufacturer's instructions. RNase Out (GIBCO, Invitrogen) was added to the RNA solutions to prevent degradation by RNase. Total RNA served as the template for cDNA preparation using the Bio-Rad one-step cDNA preparation kit. Primers were designed from rat and human cDNA sequences using Primer Express Software purchased from Applied Biosystems (Perkin-Elmer). β-Actin was used as a reference gene to normalize the relative expression of each RAS gene. The sequences of primers used are shown in Table 1. Sybergreen reactions were set up in a volume of 25 μl with ABI two-step sybergreen PCR reagents. Each reaction consisted of 12.5 μl PCR master mix, 0.05–0.30 μM of each amplification primer, and 1 μl cDNA. Each sample was run in duplicate with an initial 10-min period at 95°C to enable the reaction, followed by 40 cycles at 95°C for 15 s and 60°C for 1 min. The samples were heated to 60°C for 1 min, then to 95°C over the next minute, and finally cooled slowly from 95°C to 60°C over 20 min to collect data for the analysis of dissociation curve. Amplification data were collected by the 7700 Sequence Detector and analyzed with Sequence Detection System software. The RNA concentration in each sample was determined from the threshold cycle (CT) at which fluorescence was first detected, with the cycle number being inversely related to cDNA concentration. The fold changes in mRNA expression were calculated using the 2−ΔΔCT method (23).
Immunofluorescent labeling was carried out as described previously (34) with some modifications to determine the mucosal localization of Mas receptors. Isolated jejunal segments were rinsed with cold saline and then quickly transferred to ice-cold 4% paraformaldehyde (PFA) in 0.1 M PBS (pH 7.4) and incubated at 4°C overnight. Tissue segments were rinsed with PBS and incubated with 20% sucrose in PBS at 4°C overnight and later embedded in optimum cutting temperature medium (Tissue-Tek). Sections (6 μm) were collected on Superforst slides (Menzel-Glaser), and these were boiled in 10 mM citrate buffer for 10 min to retrieve the antigens. Sections were incubated with 1% BSA and 6% (wt/vol) normal donkey serum (NDS) (Jackson ImmunoResearch) for 1 h at room temperature to block nonspecific antibody binding. The slides were incubated overnight at 4°C with primary antibody (anti-Mas receptor rabbit polyclonal antibodies; Santa Cruz Biotechnology) (1:50) diluted in PBS with 2% NDS and 0.1% Triton X-100. After three washes with PBS, bound primary antibodies were detected by incubation with their corresponding secondary antibodies labeled with Cyc-3 (Jackson ImmunoResearch) (1:100, diluted with 0.1 M PBS containing 2% NDS) at room temperature for 1 h. Immunoreactivity was captured with a fluorescent microscope equipped with a DC480 digital camera (Leica Microsystems). Human Caco-2 cells were growth on silane-coated cover slips, incubated with ice-cold 4% PFA in 0.1 M PBS (pH 7.4) for 8 min, and washed three times. All of following steps are the same as the immunostaining procedures mentioned above starting from the incubation with 1% BSA and 6% (wt/vol) NDS.
d-Glucose and l-leucine uptake.
Uptake experiments were carried out using jejunal segments 3–4 cm in length taken from the midpoint of the section used for enterocyte isolation. Glucose and leucine uptake was measured using everted sleeves, a validated procedure for measuring solute uptake across the BBM (8, 19). In brief, isolated jejunal segments were rinsed with cold saline and everted over a glass rod. The tissue was tied securely to the rod and preincubated in gassed (95% O2-5% CO2) bicarbonate buffer (in mM: 128 NaCl, 4.7 KCl, 2.5 CaCl2, 1.2 KH2PO4, 1.2 MgSO4, and 20 NaHCO3) without glucose for 4 min at 37°C followed by 2 min using the same buffer to which 0.1 nM-5 μM ANG-(1–7) (Bachem) with or without 10–1,000 nM A-779 (Sigma-Aldrich), 0, 12.5, or 50 nM DX-600 (Bachem), 0.3 mM phlorizin (Sigma-Aldrich), or 0.1 mM phloretin (Sigma-Aldrich) was added. The tissue was then incubated with fresh buffer, with or without the same inhibitor, and with the addition of 50 mM d-glucose and 0.2 μCi/ml d-[14C]glucose (GE Healthcare) with trace amounts of 0.1 μCi/ml l-[3H]glucose (Sigma-Aldrich) to correct for nonspecific uptake. After 2 min incubation, the segments were washed rapidly with ice-cold saline containing 0.3 mM phlorizin stirring at high speed for 1 min. The tissue was removed from the rod, oven-dried, and weighed. The dried residue was incubated with Soluene-350 (Perkin-Elmer) at 60°C for 4 h. Scintillation fluid (Ultima Gold; Perkin-Elmer) was added, and counting of radioactivity was carried out. A similar procedure was used to measure the uptake of l-[3H]leucine (5 mM, 0.4 μCi/ml). The rate of glucose and leucine uptake was calculated as picomole per milligram dry weight intestine per second.
Glucose uptake by caco-2 cells.
The human colorectal adenocarcinoma epithelial cell line Caco-2 was purchased from ATCC (catalog no. HTB-37). Cells were grown at 37°C in minimum essential medium (M2279; Sigma-Aldrich) with 20% FBS, 1% nonessential amino acids (11140; GIBCO), and 1% penicillin-streptomycin and gassed with 95% air-5% CO2. Cells were subcultured at ∼80% confluency, at a cell density of between 8 × 104 and 1 × 105 cells/cm2. The medium was changed every 2 days, and cells were used 28 days after confluence. Upon reaching confluence, the cells express characteristics of enterocyte differentiation (18). Caco-2 cells grown in 5.6, 11.2, or 25 mM glucose for 28 days after confluence were rinsed with warm oxygenated bicarbonate buffer (in mM: 128 NaCl, 4.7 KCl, 2.5 CaCl2, 1.2 KH2PO4, 1.2 MgSO4, and 20 NaHCO3) with 10% FBS , 1% penicillin-streptomycin, and 1% nonessential amino acids. Cells were preincubated in gassed (95% O2-5% CO2) bicarbonate buffer containing 25 mM mannitol for 3 h at 37°C. The buffer was then replaced with fresh bicarbonate buffer containing 25 mM glucose and 0.2 μCi/ml d-[14C]glucose (GE Healthcare), with or without 1 μM ANG-(1–7), or 1 μM GF-109203X hydrochloride [a protein kinase C (PKC) inhibitor] and incubated for 1 h. In some experiments, 0.3 mM phlorizin or 0.1 mM phloretin was also present. Cells were washed quickly three times by stirring in ice-cold saline containing 0.3 mM phlorizin. The cells were digested with 0.3 M NaOH, and aliquots were added to scintillation fluid (Ultima Gold; Perkin-Elmer). Glucose uptake was calculated as disintegrations per minute per well of a six-well plate containing a monolayer of Caco-2 cells.
Measurement of ANG-(1–7) concentration in enterocytes.
ANG-(1–7) was detected using an EIA kit (catalog no. S-1330; Peninsula Laboratories, San Carlos, CA). Protein samples of jejunal enterocytes were prepared for assay using CytoBuster Protein Extraction reagents (71009-4; Novagen). The 96-well plate was read by a μQuant Plate Reader (Biotek Instruments, Winooski, VT), and calculations of ANG-(1–7) level were performed by KC Junior Software (Biotek Instruments).
Glucose tolerance tests.
Rats with STZ-induced T1DM of 2 wk duration or nondiabetic controls were fasted overnight. For oral glucose tolerance tests, animals were gastric gavaged with glucose (1 g/kg body wt) dissolved in 1 ml saline 30 min after a tail-vein injection of saline alone or ANG-(1–7) (1 mg/kg body wt) dissolved in saline. For intravenous glucose tolerance tests, glucose (1 g/kg body wt) dissolved in 1 ml saline was injected in the left jugular vein via a catheter. Tail vein blood samples were taken before glucose tolerance procedures and then 20, 40, 60, and 120 min after gavage or intravenous glucose. Blood glucose concentration was measured using test strips (Ascensia Elite; Bayer Healthcare), as described previously (5, 6). Insulin was measured using a Mercodia Insuline Elisa Kit (Uppsala, Sweden).
All results were analyzed by Prism 3.0 software. The data are expressed as means ± SE. Student's unpaired two-tailed t-test and one-way ANOVA were used to detect significant differences between two groups and three or more groups, respectively. For all comparisons, P < 0.05 was considered statistically significant. For RT-PCR, the CT value of the target gene of a sample was first corrected for the CT value of β-actin, before being statistically analyzed (23).
Enterocyte expression of ACE2, Mas receptor, and ANG-(1–7).
Western blotting of enterocyte protein revealed the presence of ACE2 and Mas receptor proteins in the jejunum of both control and 2-wk-old T1DM rats (Fig. 1, A and B). Levels of ACE2 and Mas receptor proteins in enterocytes from diabetic animals were increased by 1.59- and 1.55-fold, respectively, compared with controls. The real-time PCR analysis of mRNA expression of ACE2 and Mas receptor normalized to β-actin is shown in Fig. 1, C and D, respectively. Expression of mRNA for ACE2 and Mas receptor in diabetic jejunal enterocytes was increased by 2.73- and 2-fold, respectively, compared with values in control animals. Diabetes resulted in a 5.5-fold increase in levels of ANG-(1–7) in diabetic enterocytes compared with control values (Fig. 1E).
Localization of mas receptor in rat jejunal epithelium and Caco-2 cells.
Immunocytochemistry of jejunum revealed the presence of Mas receptor at both the BBM and basolateral membrane (BLM) along the entire jejuna villus length in tissue from both normal (Fig. 2A) and diabetic (Fig. 2B) animals. However, expression of Mas receptor was consistently higher in diabetic jejunum, and this was particularly apparent at higher magnification (Fig. 2D). Negative controls of the Mas receptor immunolabeling procedure in which primary antibodies were absent are shown in Fig. 2C. Immunocytochemistry also revealed the presence of Mas receptor at the cell membrane of Caco-2 cells grown in 5.6 mM glucose for 28 days after confluence (Fig. 2E). Expression of Mas receptor was higher in Caco-2 cells grown in 25 mM glucose (Fig. 2F), and this was particularly obvious at high magnification (Fig. 2, G and H). A negative control of the Mas receptor immunolabeling procedure in which primary antibodies were absent is shown in Fig. 2I.
Effects of ANG-(1–7), Mas receptor and ACE2 blockers, and glucose transporter inhibitors on jejunal glucose uptake.
We studied the role of the ACE2-ANG-(1–7)-Mas receptor axis in modulating jejunal glucose uptake. With the use of intestine from nondiabetic animals, mucosal exposure to ANG-(1–7) for 4 min decreased glucose uptake in a dose-dependent fashion (Fig. 3A). At the maximum concentration of ANG-(1–7) used (5,000 nM), uptake was inhibited by 32%. In contrast, the addition of A-779 (1,000 nM) or DX-600 (50 nM), blockers of Mas receptor and ACE2, respectively, to the mucosal buffer promoted glucose uptake by 36% (Fig. 3B) and 38% (Fig. 3C), respectively. The addition of A-779 (1 μM) negated the inhibitory effects of ANG-(1–7) (1 μM) on jejunal glucose uptake by nondiabetic intestine [P > 0.05, ANG-(1–7) + A-779] (Fig. 4A) vs. control. The presence of 0.3 mM phlorizin, a specific SGLT1 blocker, in mucosal fluid inhibited glucose uptake by some 76.5% (P < 0.001; Fig. 4B). Phloretin, a blocker of facilitated (GLUT-mediated) glucose transport, had a similar inhibitory action on glucose uptake (69.3% inhibition at 0.1 mM phloretin, P < 0.001). One micromolar ANG-(1–7) was without effect on glucose uptake when added to mucosal fluid in the presence of 0.3 mM phlorizin but induced an additional 22.4% decrease in glucose uptake in the presence of 0.1 mM phloretin (P < 0.001). Despite a clear action of ANG-(1–7) on glucose uptake, the peptide did not affect uptake of l-leucine (Fig. 4C).
Effects of ANG-(1–7) and glucose transporter inhibitors on jejunal glucose uptake of rats with T1DM.
TIDM promoted jejunal glucose uptake by 56% (Fig. 5A). Addition of 5 μM ANG-(1–7) inhibited the increased glucose uptake by 30.7%. The presence of 0.3 mM phlorizin in mucosal fluid inhibited glucose uptake by some 63.2% (P < 0.001; Fig. 5B). Phloretin had a similar inhibitory action on glucose uptake (40.3% inhibition at 0.1 mM phloretin, P < 0.01). Five micromolar ANG-(1–7) was without effect on glucose uptake when added to mucosal fluid with 0.3 mM phlorizin but further reduced glucose uptake in the presence of 0.1 mM phloretin (P < 0.001).
Effects of increasing glucose concentration on the expression of RAS components in Caco-2 cells.
The Caco-2 cell line was used to study the effects of glucose concentration in the growth medium on gene expression of RAS components and to further examine aspects of the ACE2-ANG-(1–7)-Mas receptor signaling pathway. To eliminate possible osmotic effects of 25 mM glucose, all buffers contained varying amounts of d-mannitol to make the final concentration of glucose and/or mannitol equal to 25 mM. mRNA expression of major RAS components was detected in Caco-2 cells and regulated by the glucose concentration in the growth medium. Thus a raised glucose concentration (25 mM) decreased the gene expression of AT1 receptor (Fig. 6A), AT2 receptor (Fig. 6B), and ACE (Fig. 6C) by 89.5, 79.6, and 80.1%, respectively. However, 25 mM glucose promoted the expression of ACE2 and Mas receptor by 264 and 193%, respectively (Fig. 6, D and E). High glucose levels in the growth medium increased gene expression of SGLT1 (Fig. 6F) and GLUT2 (Fig. 6G). Similar to jejuna enterocytes, high glucose concentrations resulted in a significantly increased level of ANG-(1–7) in Caco-2 cells compared with control values (Fig. 6H).
Effects of ANG-(1–7) and glucose transporter inhibitors on glucose uptake of Caco-2 cells.
ANG-(1–7) inhibited glucose entry into Caco-2 cells in a dose-dependent fashion (Fig. 7A). Thus 1 μM ANG-(1–7) reduced glucose uptake by 30.3%. The time course for glucose uptake by Caco-2 cells showed that an increase in incubation time from 15 to 60 min and from 1 to 3 h elevated glucose uptake by 75 and 175%, respectively (Fig. 7B). A prolonged incubation time (20 h) resulted in an eightfold increased uptake. Addition of 0.3 mM phlorizin or 0.1 mM phloretin to uptake buffer did not further decrease glucose uptake induced by ANG-(1–7) (P > 0.05) (Fig. 7C). The presence of 1 μM GF-109203X (a specific inhibitor of PKC) attenuated the inhibitory actions of 1 μM ANG-(1–7) (Fig. 7D).
Effects of ANG-(1–7) on glucose tolerance.
We examined the in vivo effects of ANG-(1–7) on oral glucose tolerance in normal and T1DM rats. Results showed that a single intravenous injection of ANG-(1–7) (1 mg/kg) improved the oral glucose tolerance of diabetic but not control animals (Fig. 8A). The effect of ANG-(1–7) was readily observable when the blood glucose data were expressed as the area under the curve (AUC, Fig. 8B). The 120-min AUC value of ANG-(1–7)-treated T1DM rats was 22.1% lower compared with untreated animals (P < 0.05). Insulin was undetectable in plasma of T1DM rats. In contrast to oral glucose tolerance data, no effect of ANG-(1–7) was noted on intravenous glucose tolerance in T1DM rats (Fig. 8C). The 120-min AUC values of ANG-(1–7)-treated and nontreated T1DM rats were not statistically significant (P > 0.05; Fig. 8D).
T1DM promotes glucose transport across the rat intestinal BBM, a response that involves increased SGLT1-mediated transport together with recruitment of GLUT2 to this membrane (8, 9, 22, 33). These events are closely linked, since enhanced SGLT1-mediated glucose transport contributes to the raised BBM levels of GLUT2 during diabetes, a process mediated by the PKC signaling pathway (21). Expression of SGLT1 is highly regulated at both local and systemic levels. Indeed, many factors have been postulated to be responsible for the raised SGLT1-mediated transport during diabetes (10). Our previous work has assessed the involvement of the local ACE-ANG II-AT1 receptor axis in the control of jejunal glucose uptake. We showed that all components of this system are expressed in jejunal enterocytes and that ANG II, acting via AT1 receptors, rapidly suppressed SGLT1-mediated glucose uptake (34). Furthermore, TIDM reduced expression of ACE and AT1 receptor at the enterocyte BBM, and we proposed that this might contribute to upregulated SGLT1-mediated glucose transport in diabetes (33).
A growing body of evidence suggests that the ACE2-ANG-(1–7)-Mas receptor axis opposes the actions of the ACE-ANG II-AT1 receptor system in many tissues, including the heart, liver, and kidney (2). It has been shown that expression of ACE2 protein is higher in the kidneys of mice with T1DM (36), and ACE2 has been detected in small intestine (31). We thus decided to investigate the influence of this system on glucose uptake across the enterocyte BBM and to assess the effects of T1DM on jejunal expression of the ACE2-ANG-(1–7)-Mas receptor axis. Our findings indicate that enterocyte levels of ANG-(1–7), ACE2, and Mas receptor are higher in T1DM and that ANG-(1–7) exerts a dose-dependent inhibitory effect on Mas receptor-mediated jejunal glucose uptake.
Our previous finding of reduced expression of ACE and AT1 receptor in jejunal enterocytes during T1DM would decrease enterocyte production of ANG II (33), and this may explain our finding of raised levels of ACE2 and ANG-(1–7) in diabetic enterocytes, since ANG II inhibits the formation of ACE2 in other cell types (14, 15). Immunodetection showed the localization of Mas receptor along both the BBM and BLM of the entire jejunal villus axis, a profile similar to that seen for AT1 and AT2 receptors (33). Immunoreactivity in tissue from diabetic animals was more intense than nondiabetic animals, a finding in keeping with protein and mRNA expression data. Previous studies using a wide range of cell systems indicate a complex regulation of the signaling mechanisms that determine the actions of ANG II and ANG-(1–7). ANG-(1–7) can act as an endogenous allosteric modulator of AT1 or AT2 receptors by changing their binding and functional responses (4), whereas ANG II is able to modulate the activity of the ACE2 system. In enterocytes, upregulated expression of ANG-(1–7) may have a role in balancing the adverse effects caused by downregulation of the ACE-ANG II-AT1 receptor axis. There is much evidence in other tissues that this balancing process opposes disease progression. Thus, raised ACE2 expression in prehypertensive subjects moderates progression of this disease (20), ANG-(1–7) administration to rats reversed chemically induced glomerulonephritis (37), administration of recombinant ACE2 to mice reduces the pressor actions of ANG II while increasing plasma levels of ANG-(1–7) (35), and treatment of rats with an ANG-(1–7) antagonist or an ACE2 inhibitor worsened the course of two-kidney, one-clip hypertension (3). Furthermore, increased expression of ACE2 occurs in glomeruli and peritubular capillary endothelium of patients with primary and secondary renal disease, including transplanted kidneys (25), whereas mice lacking Mas receptor display glucose intolerance, lowered insulin sensitivity, and reduced insulin-stimulated glucose uptake and GLUT4 expression in adipose tissue (28). Taken together, these studies indicate that the ACE2-ANG-(1–7)-Mas receptor axis has an important protective role at the local level.
In contrast to the situation in other tissues, our present work indicates that ANG II and ANG-(1–7) peptides do not have opposing actions on enterocyte glucose uptake. Indeed, the action of both peptides is to suppress SGLT1-mediated uptake. However, differential changes in enterocyte expression of the ACE-ANGII-AT1 receptor (33, 34) and ACE2-ANG-(1–7)-Mas receptor systems in response to diabetes, and perhaps other diseases of glucose metabolism, produce opposing actions on glucose uptake. The use of phlorizin and phloretin to block SGLT1- and GLUT-mediated uptake, respectively, in our present study revealed that ANG-(1–7), like ANG II (34), blocked SGLT1-mediated glucose uptake. Caco-2 cells grown in medium containing a glucose concentration similar to that seen in blood of T1DM rats (8, 9) showed similar changes in the expression of RAS components to those induced by T1DM in rat jejunum (33). Caco-2 cells are therefore a potentially good model to study downstream signaling pathways that culminate in altered expression of glucose transporters following ANG-(1–7) binding to Mas receptor. The use of the specific blockers GF-109203X hydrochloride and A-779 in Caco-2 cells and rat jejunum suggests that ANG-(1–7) achieves its action in enterocytes via the binding to Mas receptor and subsequent activation of the PKC signaling pathway. However, the precise relationship between the Mas receptor, PKC activation, and reduced SGLT-mediated glucose transport in jejunum is unknown at the present time. The inhibition of BBM glucose uptake by ANG-(1–7) seen using both jejunum and Caco-2 cells may have implications for control of postprandial glycemia in diabetes mellitus. In this context, it was of importance that treatment of diabetic rats with ANG-(1–7) significantly improved tolerance of an oral but not an intravenous glucose load, and this strongly suggests an action of the peptide on intestinal glucose transport. Taken together, our findings imply that ANG-(1–7) is able to inhibit glucose uptake by an action from either the luminal or blood side of the jejunal epithelium, and this may be important in the search for novel therapies to ameliorate the metabolic repercussions of postprandial hyperglycemia in diabetics.
In summary, we have shown for the first time the expression of the components of the ACE2-ANG-(1–7)-Mas receptor system in jejunal enterocytes and the involvement of ANG-(1–7) in the control of SGLT1-mediated glucose transport across the intestinal BBM. It is therefore clear that the RAS provides multiple pathways for the local control of enterocyte glucose transport. Upregulated expression of the ACE2-ANG-(1–7)-Mas receptor axis during T1DM may counterbalance the higher glucose uptake induced by downregulation of the ACE-ANG II-AT1 receptor axis in this condition. Our study raises the possibility for the jejunal ACE2-ANG-(1–7)-Mas receptor axis to be targeted therapeutically in the search for novel treatments for glycemic regulation in diabetic patients.
The work described in this paper was fully supported by a grant from the Research Grants Council of the Hong Kong Special Administrative Region, China (Project no. CUHK470709), awarded to P. S. Leung.
No conflicts of interest, financial or otherwise, are declared by the authors.
T.P.W. and K.Y.H. performed experiments; T.P.W., K.Y.H., and P.S.L. analyzed data; T.P.W. and P.S.L. prepared figures; T.P.W. and P.S.L. drafted manuscript; E.K.W.N., E.S.D., and P.S.L. interpreted results of experiments; E.S.D. and P.S.L. edited and revised manuscript; P.S.L. conception and design of research; P.S.L. approved final version of manuscript.
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