Angiotensin II Induces Interleukin-1β–Mediated Islet Inflammation and β-Cell Dysfunction Independently of Vasoconstrictive Effects

Obesity and type 2 diabetes are related to hypertension and to increased activation of the renin-angiotensin system (RAS) (1–3). Multiple trials have shown that RAS blockade reduces the incidence of new-onset type 2 diabetes in high-risk populations (4). On the basis of several meta-analyses, this reduction ranges between 22% and 30% (5,6). In addition, in different animals models of type 2 diabetes, treatment with either angiotensin-receptor blockers or ACE inhibitors improves glucose tolerance and β-cell function (2,7–10). All of this suggests a role for angiotensin II in the development of type 2 diabetes.

The RAS is classically known as a systemic hormonal system regulating blood pressure, fluid balance, and electrolyte absorption (11). Finding a local RAS in various tissues and organs such as brain, kidney (12), heart (13), liver, and adipose tissue (14) has expanded its role to diverse physiological functions in addition to its effects on circulation. All key components of the RAS also have been localized to the endocrine pancreas, including the precursor angiotensinogen and the angiotensin II type 1 receptor (15,16). Furthermore, obesity and hyperglycemia increases the expression of the local RAS in pancreatic islets (17), adipose tissue (18), and skeletal muscle.

Several hypotheses of how RAS activation might contribute to the development of diabetes and why its blockade could be protective have been suggested. In diabetic animal models, angiotensin II leads to decreased blood flow in insulin target tissues and pancreatic islets, which results in reduced insulin and glucose delivery (19–21). In skeletal muscle angiotensin II interferes with glucose uptake by decreasing GLUT4 translocation to the plasma membrane (22) and induces insulin resistance (23). RAS inhibition prevents these effects in vivo, resulting in increased glucose tolerance and improved islet function (2,8,9,24). However, whether the amelioration of metabolic parameters is a consequence of normalization of vasoconstriction or due to inhibition of local RAS is unclear. Studies of isolated islets treated with angiotensin II or its blockers point to a possible role of activated local RAS or direct angiotensin II effects on impaired insulin secretion (15,17,25).

It has been recognized more recently that in type 2 diabetes chronic inflammation is involved in the dysfunction of pancreatic islets (26,27). Increased numbers of immune cells were observed in pancreatic islets of animals fed a high-fat diet (HFD) and of patients with type 2 diabetes (28). There is increasing evidence that islet inflammation is mediated by an imbalance of interleukin (IL)-1β and its naturally occurring antagonist IL-1 receptor antagonist (IL-1Ra). This contributes to the formation of insulitis by recruiting and activating IL-1β–producing macrophages. Treatment of type 2 diabetes or obese patients with anakinra, the recombinant form of IL-1Ra, or specific anti–IL-1β antibodies improved glycemia and β-cell function and reduced circulating inflammatory indicators (29–34).

Some observations point to a possible proinflammatory role for angiotensin II (35,36). It triggers inflammatory processes in the kidney (37) and induces the chemokine MCP-1 in pancreatic cancer cells (38). In blood mononuclear cells, ACE inhibitors suppress IL-1 and tumor necrosis factor (TNF) synthesis (39). Furthermore, in clinical trials treatment with angiotensin II receptor antagonists reduced the proinflammatory markers TNF-α, IL-6, and CRP (40,41), as well as MCP-1, in the circulation of patients with cardiovascular diseases (42). Similarly, in HFD-fed mice, decreased serum concentrations of interferon-γ and MCP-1 and diminished proinflammatory gene expression in pancreatic islets were observed with angiotensin II receptor antagonists or ACE inhibitors (24,43).

All of this suggests a role for angiotensin II in the development of inflammation and type 2 diabetes. However, a possible direct effect of angiotensin II on metabolism and insulin secretion independent of changes in blood pressure and the underlying pathway remain to be investigated.

Human islets were isolated from pancreata of cadaver organ donors in the islet transplantation centers in Lille (France) and Geneva (Switzerland), in accordance with the local institutional ethical committee. Human islets were provided by the islets for research distribution program through the European Consortium for Islet Transplantation, under the supervision of the Juvenile Diabetes Research Foundation (31-2012-783). Islets were cultured in CMRL-1066 medium containing 5 mmol/L glucose, 100 units/mL penicillin, 100 µg/mL streptomycin, 2 mmol/L glutamax, and 10% FCS (Invitrogen, Basel, Switzerland) on extracellular matrix–coated 24-well plates (Novamed Ltd, Jerusalem, Israel) in a humid environment containing 5% CO₂. Islets were treated for 96 h (media were renewed after 48 h) with or without 1 μmol/L angiotensin II (A9525; Sigma Aldrich, Switzerland), 1 µg/mL IL-1Ra (Kineret; Amgen, Thousand Oaks, CA), and/or 10 μmol/L IKK-2 inhibitor (SC-514; Merck, Darmstadt, Germany). Culture supernatants were collected and islets were used for RNA extraction or glucose-stimulated insulin secretion experiments.

To isolate mouse islets, pancreata were perfused with a collagenase solution (Worthington, Lakewood, NJ) and digested in the same solution at 37°C, followed by filtration through 500- and 70-µm cell strainers. Islets were cultured in RPMI-1640 medium containing 11.1 mmol/L glucose, 100 units/mL penicillin, 100 µg/mL streptomycin, 2 mmol/L glutamax, 50 µg/mL gentamicin, 1:1,000 Fungizone (Gibco), and 10% FCS. Islets were directly collected for RNA extractions, were cultured for 36 h on extracellular matrix–coated 24-well plates and treated for 24 h with 1 μmol/L angiotensin II before RNA extraction and protein measurements in the supernatant, or were used for glucose-stimulated insulin secretion experiments (48 h treatment with 1 μmol/L angiotensin II).

INS-1E cells (44) were cultured in RPMI-1640 medium containing 11.1 mmol/L glucose, 100 units/mL penicillin, 100 µg/mL streptomycin, 2 mmol/L glutamax, 10% FCS, 10 mmol/L HEPES, 1 mmol/L sodium pyruvate, and 50 μmol/L 2-mercaptoethanol. For experiments, 100,000 cells/well were seeded in 24-well plates. Cells were treated with or without 1 μmol/L angiotensin II and/or 10 μmol/L IKK-2 inhibitor for 72 h before RNA extraction. For glucose-stimulated insulin secretion experiments, cells were treated 96 h with 1 μmol/L angiotensin II. Apoptosis rates were measured with the Cell Death Detection Elisa Kit (Roche Diagnostics, Switzerland) according to the manufacturers’ instructions.

For in vitro or ex vivo glucose-stimulated insulin secretion experiments, islets or INS-1E cells were seeded for 2 days in quadruplicate. After treatment with angiotensin II, supernatants were collected and stored at −20°C (chronic insulin release). Cells were preincubated for 30 min in modified Krebs-Ringer bicarbonate buffer (mKRBB; 115 mmol/L sodium chloride, 4.7 mmol/L potassium chloride, 2.6 mmol/L calcium chloride dihydrate, 1.2 mmol/L monopotassium sulfate, 1.2 mmol/L magnesium sulfate heptahydrate, 10 mmol/L HEPES, and 0.5% bovine serum albumin [pH 7.4]) containing 2.8 mmol/L glucose. mKRBB was then replaced by mKRBB 2.8 mmol/L glucose for 1 h (basal insulin release), followed by 1 h in mKRBB 16.7 mmol/L glucose (stimulated insulin release). Islets or INS-1E cells were extracted with 0.18 N hydrogen chloride in 70% ethanol to determine insulin content. Insulin concentrations were determined using human or mouse insulin ultrasensitive ELISA (Mercodia, Uppsala, Sweden) or mouse/rat insulin kit (Mesoscale Discovery, Rockville, MD). Stimulatory index was determined as ratio of insulin secretion at 16.7 to that at 2.8 mmol/L glucose/hour.

Oxygen consumption rates were determined using the Seahorse extracellular flux analyzer XF^e96 (Seahorse Bioscience). INS-1E cells (145,000 cells/well) were seeded on poly-d-lysine-treated Seahorse 96-well microplates in 175 µL INS-1E medium/well 2 days before the experiment. Medium was changed and cells were treated with 1 μmol/L angiotensin 24 h before the experiment. At the day of the assay, cells were preincubated in unbuffered assay medium (RPMI-1640 [R6504; Sigma] supplemented with 11.1 mmol/L glucose) for 1.5 h at 37°C in air. To test ATP turnover, maximal respiratory capacity, and nonmitochondrial respiration of the cells, 1 μmol/L oligomycin, 2 μmol/L of the uncoupler carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone, and 1 μmol/L rotenone were successively injected.

To determine β-cell apoptosis, human islets were dispersed with trypsin-EDTA (Invitrogen) for 6 min at 37°C. Single cells were cultured and treated with angiotensin II for 4 days (new medium and treatment were added after 2 days). Subsequently, the cells were fixed in 4% paraformaldehyde for 30 min, followed by incubation in 0.5% Triton-X-100 for 4 min. Cells were triple stained with the terminal deoxynucleotidyl TUNEL technique (In Situ Cell Death Detection Kit; Roche Diagnostics, Switzerland), a polyclonal antibody against insulin (Dako, Denmark), and nuclear DAPI (Sigma).

Total RNA of isolated human or mouse islets and INS-1E cells was extracted using the Nucleo Spin RNA II Kit (Machery Nagel, Düren, Germany). cDNA was prepared with random hexamers and Superscript II (Invitrogen). For quantitative PCR, the real-time PCR system 7500 (Applied Biosystems) and the following TaqMan assays were used: human IL-1β, Hs00174097_m1; IL-6, Hs00174131_m1; IL-8, Hs00174103_m1; 18s, Hs99999901_s1; MCP-1, Hs00234140_m1; rat CXCL1, Rn00578225_m1; and mouse GAPDH, Mm99999915_g1; IL-6: Mm00446190_m1. Gene expression was analyzed using the comparative 2^-ΔΔCT method.

Human or mouse IL-6 concentrations in islet culture supernatants or plasma were assayed using Luminex technology (Millipore, Billerica, MA) or Mesoscale kits (Mesoscale Discovery).

Formalin-fixed pancreata were embedded in paraffin and blocks were cut at 50-µm intervals. Tissue sections were deparaffinized, rehydrated, and stained with rat antimouse CD45 (BD Bioscience, New Jersey), rat antimouse F4/80 (Cedarlane, Burlington, ON), or rabbit antimouse CD3 antibody (Abcam, Cambridge, U.K.). To determine immune cell infiltration, two to five slides per mouse were stained; CD45⁺ cells were counted under a microscope (Olympus BX63) and the percentages of affected islets per mouse were calculated. In the angiotensin II and hydralazine group, 862 islets were counted and 28 islets were affected (10 mice, pooled from three experiments); in the angiotensin II and hydralazine plus anti–IL-1β antibody group, 458 islets were counted (four mice; eight islets were affected).

Male C57BL/6N mice were obtained from Charles River Laboratories (Sulzfeld, Germany) at 4 weeks of age. For the first three experiments, a total of 78 mice were used; in the fourth experiment 17 mice were used. They were fed an HFD (D12331; Research Diets, New Brunswick, NJ), and after 12 weeks they were subcutaneously implanted with osmotic mini pumps (Alzet 2004; Durect Corp., Cupertino, CA) releasing either angiotensin II (1 µg/kg/min; A9525; Sigma) or saline for 4 weeks. Thirty-nine animals in the first three experiments and all 17 animals in the fourth experiment received hydralazine (H1753; Sigma) in their drinking water at a concentration of 250 mg/L. Animals used in the fourth experiment were injected subcutaneously once per week for 4 weeks with either saline (five mice), control antibodies (anti-cyclosporine-A; four mice), or mouse anti–IL-1β antibodies at a concentration of 10 µg/g (eight mice). The anti–IL-1β antibody is a mouse antibody with the same specificity as canakinumab (45). Antibodies were kindly provided by Novartis (Basel, Switzerland). Surgery was done in a specific pathogen–free environment, animals were anesthetized with Ketalar (65 mg/kg) and Xylasol (13 mg/kg) by intraperitoneal injection. To prepare for pump implantation, an air pocket was created under the skin. The sterile pumps (filled with saline or angiotensin II) were inserted in the pocket and the wound was closed with two clips. After the first signs of waking, an analgesic (Temgesic, 0.05 mg/kg) was injected subcutaneously. After 24 h and, if necessary, 48 h, another injection of painkiller was given. Wound healing and the health state of every mouse was observed and recorded on a score sheet every day for 1 week. All animals were housed singly in cages in a temperature-controlled room with a 12-h light/12-h dark cycle and were allowed free access to food and water, according to Swiss veterinary law and institutional guidelines. After 4 weeks of treatment mice were used for glucose/insulin tolerance tests. After death, heart blood was collected and islets were isolated for RNA expression or ex vivo glucose-stimulated insulin secretion experiments, or pancreata were taken for histology.

Blood pressure was measured with a tail-cuff system (Visitech Systems, Apex, NC) in six to eight mice per group. Mice were first habituated to the tail-cuff system for 5 days to avoid stress artifacts caused by the demanding procedure. Every day they were put into the system and the tails were fixed in a cuff. After training the mice, data were acquired on the following 3–5 days. While discarding the first 10 measurements, the subsequent 10 measurements were averaged and reported.

For intraperitoneal glucose tolerance tests, mice were fasted for 6 h in the morning and injected intraperitoneally with 2 g glucose/kg body weight. Blood samples for glucose measurements were obtained at 0, 15, 30, 60, 90, and 120 min using a glucometer (Freestyle; Abbott Diabetes Care Inc., Alameda, CA) and at 0, 15, and 30 min for measurement of plasma insulin concentrations using an insulin ELISA (Mercodia). For intraperitoneal insulin tolerance tests, mice were fasted 3 h in the morning before administration of 1 unit/kg insulin (Novo Nordisk, Bagsvaerd, Denmark), and blood glucose was measured at 0, 15, 30, 60, and 90 min.

Statistical analysis was performed using GraphPad Prism 5 (Graphpad Software Inc., San Diego, CA). Data are presented as mean ± SEM and were analyzed using one- or two-way ANOVA or unpaired Student t test. Differences were considered statistically significant when P < 0.05.

To investigate the effect of angiotensin II on β-cell function and survival in vitro, we exposed isolated human and mouse islets to 1 μmol/L angiotensin II for 48 to 96 h. This treatment resulted in a slight increase of basal and a decrease of glucose-stimulated insulin secretion (Fig. 1A and C), leading to a significant decrease of the stimulatory index overall compared with control islets (Fig. 1B and D). To discriminate whether the observed effects in whole islets are the result of (partly) direct signaling of angiotensin II on β-cells or indirect via paracrine actions, we used the β-cell line INS-1E and observed significantly higher basal insulin secretion and reduction of the stimulatory index following exposure to angiotensin II (Fig. 1E and F), as well as diminished mitochondrial respiratory capacity (Fig. 1G). Staining of treated and untreated single human islet cells with the TUNEL assay revealed a threefold increase of angiotensin II–induced β-cell apoptosis (Fig. 1H and I). Similarly, exposure of INS-1E cells to 1 μmol/L angiotensin II for 3 days induced a 1.3-fold increase of apoptosis (data not shown).

Next we questioned whether angiotensin II has proinflammatory effects in isolated human and mouse islets. Angiotensin II increased the gene expression of several cytokines, including IL-1β, MCP-1, IL-8, and IL-6 (Fig. 2A–D). This led to elevated IL-6 protein release into the culture supernatant of human islets (Fig. 2E). Similarly, IL-6 expression and release was stimulated by angiotensin II in mouse islets (Fig. 2F and G). TNF was not significantly changed by angiotensin II, whereas interferon-γ was not detectable (not shown). In the β-cell line INS-1E induction of the chemokine CXCL1 (functional homolog to human IL-8 [CXCL8]) (Fig. 2H) could be detected. Angiotensin II–induced increase in IL-1β and IL-6 in human islets and increased CXCL1 gene expression in INS-1E cells was mediated by nuclear factor (NF)-κB; it was fully prevented by the addition of an IκB-kinase-2 inhibitor (Fig. 3A–C). Finally, blocking IL-1 signaling with IL-1Ra prevented the angiotensin II–induced increase in IL-6 gene expression (Fig. 3D) and release (Fig. 3E), demonstrating that elevated IL-6 concentrations depend on IL-1 signaling.

To investigate the role of angiotensin II in the development of impaired insulin secretion in the context of type 2 diabetes, mice were fed an HFD for 12 weeks; then, osmotic pumps releasing either saline or angiotensin II for 4 weeks were implanted. Exposure to angiotensin II leads to vasoconstriction and hypertension, which may affect glucose tolerance because of altered glucose and insulin delivery to the insulin-sensitive tissues. Therefore, half of the animals also received the vasodilator hydralazine in the drinking water (Fig. 4A). As shown in Fig. 4B, vasoconstrictive effects of angiotensin II were present and hydralazine application reduced the angiotensin II–induced increase in mean arterial pressure. Mean body weight did not change in mice receiving angiotensin II or hydralazine alone compared with the saline group and was reduced in mice treated with the combination of the two compared with both the saline and the angiotensin II groups alone (Fig. 4C).

To assess glucose metabolism, we performed intraperitoneal glucose tolerance tests after 16 weeks on an HFD and angiotensin II treatment during the last 4 weeks. In mice with angiotensin II treatment alone, blood glucose as well as plasma insulin concentrations increased only marginally after intraperitoneal administration of glucose because vasoconstriction leads to reduced glucose absorbance. This is supported by an intravenous glucose tolerance experiment where glucose tolerance was reduced in angiotensin II–infused animals (data not shown). As shown in Fig. 4D and E, compared with saline, hydralazine improved glucose tolerance and insulin secretion in mice fed an HFD. In contrast, treatment with angiotensin II plus hydralazine led to highly impaired glucose tolerance along with a complete lack of glucose-stimulated insulin secretion (Fig. 4D and E). To assess whether increased insulin resistance contributed to impaired glucose metabolism in the combined angiotensin II and hydralazine group, we performed insulin tolerance tests (Fig. 4F). Even in the presence of angiotensin II, hydralazine improved insulin sensitivity compared with the saline group (Fig. 4F). Therefore, the glucose intolerance observed with angiotensin II and hydralazine treatment is a result of β-cell failure and is not caused by impaired insulin sensitivity.

After 16 weeks of HFD feeding and 4 weeks of treatment with angiotensin II and hydralazine, IL-6 plasma concentrations were increased in treated mice compared with control animals or mice receiving hydralazine alone (Fig. 4G). In contrast to the in vivo situation, ex vivo glucose-stimulated insulin secretion assays revealed no differences in basal and glucose-stimulated insulin concentrations in the various groups (Fig. 4H). However, chronic insulin release over 36 h into the culture medium containing 11.1 mmol/L glucose was significantly elevated in islets isolated from the angiotensin II and hydralazine group (Fig. 4I), along with increased expression of IL-6 (Fig. 4J), but the latter did not reach statistical significance.

Since angiotensin II induces IL-1β in vitro, leading to IL-6 expression, we next investigated whether the angiotensin II–induced impaired insulin secretion could also be mediated by IL-1β in vivo. For this, mice fed an HFD for 16 weeks were treated with angiotensin II and hydralazine for the last 4 weeks, as in the previous experiments. In addition, they were injected with either specific antibodies against IL-1β or with saline or control antibodies (anti-cyclosporine-A), as shown in Fig. 5A. Glucose tolerance tests revealed that inhibiting IL-1β improved glycemia and restored insulin secretion compared with control groups (glucose tolerance tests of animals treated with saline and control antibodies were identical and therefore pooled; Fig. 5B and C). The improvement in metabolism was not caused by changes in insulin sensitivity (Fig. 5D). Ex vivo, islets isolated from mice treated with anti–IL-1β antibodies had significantly lower basal and higher glucose-stimulated insulin secretion compared with islets from animals treated with saline or control antibodies, resulting in an improved stimulatory index (Fig. 5E and F).

Immunohistochemical stainings of pancreata with the panimmune cell marker CD45 revealed differences in immune cell infiltration (patterns of insulitis; examples are shown in Fig. 5H–K) in islets of mice treated with anti–IL-1β antibodies compared with the angiotensin II and hydralazine control group. The number of islets with mild forms of infiltration, as well as the total percentage of affected islets per mouse, were reduced in mice treated with angiotensin II, hydralazine, and anti–IL-1β antibodies compared with angiotensin II and hydralazine alone (Fig. 5G). Immunohistochemical stainings of CD3 and F4/80 indicated that infiltrating cells were mostly T cells and not macrophages (data not shown).

In this study we demonstrate that angiotensin II deteriorates glucose metabolism through deleterious effects on pancreatic β-cell mitochondrial function and insulin secretion. This effect involves IL-1β– and NF-κB–mediated inflammation and apoptosis and is independent of changes in blood pressure and insulin sensitivity.

In vivo infusion of angiotensin II for 4 weeks completely abolished glucose-stimulated insulin secretion. Surprisingly, after isolation and culturing, islets from angiotensin II–treated mice showed similar insulin secretion upon glucose stimulation as islets from saline-infused mice. Several possibilities can explain this finding: Angiotensin II–induced damage is reversible, and the islets had recovered between isolation and the insulin secretion assay. Alternatively, the in vivo deleterious effect of angiotensin II may be mediated via immune cells, which are lost following islet isolation. This would explain the discrepancy between the mild, direct in vitro effects of angiotensin II (Fig. 1) on insulin secretion compared with the strong in vivo impairment of insulin secretion (Fig. 4). A similar discrepancy between in vivo and in vitro islet function was observed in other animal models of diabetes displaying islet inflammation (46,47).

Our in vitro observations that angiotensin II impairs glucose-stimulated insulin secretion are in line with previous findings in mouse islets showing that treatment with angiotensin II dose-dependently reduced insulin secretion and synthesis (15). The impairment of the glucose-stimulated secretory function goes together with increased basal insulin secretion in all our in vitro models (Fig. 1). This may be because of the deleterious effects of angiotensin II on β-cells, forcing inadequate continuous release of insulin with consecutive decreased responsiveness to an acute glucose challenge. The data from our study are in apparent contrast to some older studies showing that angiotensin II induces insulin secretion in the absence of glucose stimulation (20). In the presence of large amounts of glucose, however, angiotensin II diminished mitochondrial function and insulin secretion. In MIN6 β-cells, 1 h of angiotensin II treatment potentiated glucose-stimulated insulin secretion (48). Thus acute exposure to angiotensin II promotes insulin secretion, whereas chronic treatment in the presence of elevated glucose concentrations is deleterious. This possibly reflects context-dependent effects of angiotensin II, which may be physiologic or pathologic. In line with this thinking, short-term and low-dose IL-1β stimulates insulin secretion and β-cell survival, whereas prolonged exposure is deleterious (49,50).

Hydralazine has commonly been used to reverse the hypertensive effect of angiotensin II (51). In accordance with these findings, blood pressure measurements in our in vivo experiments revealed vasoconstrictive effects of angiotensin II and the ability of hydralazine to reverse them. Since our aim was to investigate the effects of angiotensin II independent of vasoconstriction, in vivo experiments with animals that received angiotensin II together with hydralazine and the respective controls were performed. Mice treated with angiotensin II and hydralazine exhibited lower body weights compared with saline-treated animals, as previously described (52,53). Despite the lower body weight, the animals exhibited impaired glucose tolerance. Both lower body weight and glucose intolerance can be explained by a lack of insulin. This strengthens the assumption that angiotensin II impairs insulin secretion, which was not always observed when hypertension was not corrected (22,48).

Experiments with dispersed human islets revealed an increase in β-cell apoptosis after treatment with angiotensin II. Because cultured human islets grow on multiple layers, which makes the identification of β-cells difficult, whole-islet stainings were limited by technical issues. Although we acknowledge that single β-cells may behave differently than intact islets, at least we are confident that the apoptotic process occurred in β-cells. Corroboratively, we see similar proapoptotic effects of angiotensin II in the β-cell line INS-1E.

Angiotensin II induced proinflammatory cytokines in human islets, including IL-6, IL-1β, IL-8, and MCP-1, reflecting a proinflammatory state in general. We focused on IL-6 as an easily detectable marker of inflammation, not meaning that it is causal for all the effects. Angiotensin II–induced inflammation does not seem to be species dependent since mouse islets showed similar results. Moreover, our data with the pure β-cell line INS-1E indicate that there is a direct proinflammatory effect of angiotensin II on β-cells, although cell lines express a much smaller variety of cytokines.

Cytokine induction seems to be mediated via NF-κB because treatment with IκB-kinase-2 inhibitor fully prevented the angiotensin II–induced upregulation of IL-1β and IL-6 in human islets and CXCL1 in INS-1E cells.

Importantly, by using IL-1Ra, we show that the angiotensin II–mediated induction of IL-6 in human islets depends on IL-1β signaling. IL-1β is a master proinflammatory mediator involved in the development of type 2 diabetes (26); therefore its inhibition is currently in clinical development for the treatment of diabetes. In this study we show that IL-1β also mediates the deleterious effects of angiotensin II on insulin secretion and glucose homeostasis, adding to the rationale for the use of IL-1 antagonism in the treatment of the metabolic syndrome.

Taken together, our results show that chronically elevated angiotensin II concentrations induce β-cell dysfunction in vitro and in vivo, independently of its effects on blood pressure. This effect seems to be mediated by a proinflammatory response via the IL-1β/NF-κB pathway. Therefore, some of the protective effects of ACE inhibitors observed in patients with prediabetes and diabetes could be due to the prevention of angiotensin II–induced islet inflammation.

Acknowledgments. The authors thank their technicians Marcela Borsigova, Kaethi Dembinski, and Richard Prazak for excellent technical assistance.

Funding. This work was financially supported by grants from the Swiss National Science Foundation (310030-146840/1 and 32003B-130008/1 to M.Y.D. and 31003A-144112/1 to A.W.J.).

Duality of Interest. M.Y.D. is listed as the inventor on a patent (WO6709) filed in 2003 for the use of an IL-1 receptor antagonist for the treatment of, or prophylaxis against, type 2 diabetes. No other conflicts of interest relevant to this article have been reported.

Author Contributions. N.S.S., A.W.J., M.B.-S., and M.Y.D. designed the study. N.S.S., C.T., Y.P., and K.K. performed and analyzed experiments. N.S.S., C.T., M.B.-S., and M.Y.D. wrote the manuscript. E.D., S.T., and K.T. helped with the experiments. B.B., F.P., and J.K.-C. isolated human islets. M.Y.D. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.