Chronic palmitate exposure impairs glucose-stimulated insulin secretion and other aspects of β-cell function, but the underlying mechanisms are not known. Using various live-cell fluorescence imaging approaches, we show here that long-term palmitate treatment influences cAMP signaling in pancreatic β-cells. Glucose stimulation of mouse and human β-cells induced oscillations of the subplasma-membrane cAMP concentration, but after 48 h exposure to palmitate, most β-cells failed to increase cAMP in response to glucose. In contrast, GLP-1–triggered cAMP formation and glucose- and depolarization-induced increases in cytoplasmic Ca2+ concentration were unaffected by the fatty acid treatment. Insulin secretion from control β-cells was pulsatile, but the response deteriorated after long-term palmitate exposure. Palmitate-treated mouse islets showed reduced expression of adenylyl cyclase 9, and knockdown of this protein in insulinoma cells reduced the glucose-stimulated cAMP response and insulin secretion. We conclude that impaired glucose-induced generation of cAMP is an important determinant of defective insulin secretion after chronic palmitate exposure.
There is a strong link between type 2 diabetes and obesity, and the disease is often associated with increased plasma free-fatty-acid concentrations (1,2). While acute exposure of insulin-secreting β-cells to saturated fatty acids, such as palmitate, stimulates insulin secretion (3,4), long-term palmitate exposure is associated with β-cell dysfunction, including reduction of glucose-stimulated insulin secretion (5–9). The mechanisms by which chronic palmitate exposure impairs insulin secretion are incompletely understood but probably involve multiple effects at different levels (9). For example, palmitate has been reported to cause alterations in β-cell metabolism (10) and ion channel activity (11). The lipid has also been found to reduce insulin biosynthesis and to induce mitochondrial uncoupling, ceramide synthesis, and endoplasmic reticulum stress, eventually leading to apoptotic cell death (7,12–15). However, reduced insulin secretion after long-term exposure to fatty acids has been reported to occur before apparent signs of increased β-cell death, alterations of insulin synthesis, glucose metabolism, ATP-sensitive K+ channel (KATP channel) regulation, or Ca2+ signaling, suggesting that the secretion defect occurs at a late step in the exocytosis process (16), not as a result of decreased release competence of the secretory vesicles, but because they become uncoupled from the sites of Ca2+ entry (17).
The depolarization-triggered entry of Ca2+ through L-type voltage-dependent Ca2+ channels is the final step in a chain of events by which glucose triggers insulin secretion (18,19). Apart from the triggering role of Ca2+, insulin secretion is markedly amplified by cAMP. While cAMP is known to mediate the insulinotropic effects of, e.g., GLP-1 and glucose-dependent insulinotropic polypeptide (20), accumulating evidence indicates that cAMP elevation is also required for a normal insulin secretory response to glucose alone (21,22). In the current study, we used various live-cell imaging techniques to investigate whether the reduced insulin response to glucose after chronic exposure to palmitate is associated with changes of β-cell cAMP signaling.
Research Design and Methods
Adrenaline, 8-Br-cAMP, forskolin, GLP-1 (7–36) amide (GLP-1), HEPES, 2-mercaptoethanol, poly-L-lysine, palmitate, 2′,5′-dideoxyadenosine, and insulin were from Sigma (St. Louis, MO) and Lipofectamine 2000, DMEM, trypsin, penicillin, streptomycin, glutamine, and FCS were from Life Technologies (Carlsbad, CA). Plasmids and adenoviral vectors encoding a translocation biosensor for the cAMP concentration beneath the plasma membrane ([cAMP]pm) have previously been described (23). The sensor consists of a truncated and membrane-anchored protein kinase A (PKA) regulatory RIIβ subunit tagged with CFP and a PKA catalytic Cα subunit tagged with YFP (24). Insulin secretion dynamics was monitored using the PtdIns(3,4,5)P3 translocation reporter GFP4-Grp1 (21).
Islet and Cell Culture, Transfection, and Palmitate Treatment
Islets of Langerhans were isolated from C57BL/6J female mice as previously described (23). All animal handling procedures were approved by the local animal ethical committee. Human pancreatic islets from nine normoglycemic cadaveric donors (aged 47–76 years) were generously provided by the Nordic Network for Clinical Islet Transplantation. All experiments with human islets were approved by the Uppsala human ethical committee. The islets were isolated with semiautomated digestion filtration, purified on a continuous density gradient in a refrigerated cell processor (COBE 2991; COBE Blood Component Technology, Lakewood, CO) (25) and kept for 2–5 days at 37°C in an atmosphere of 5% CO2 in CMRL 1066 culture medium (Mediatech, Herndon, VA) containing 5.5 mmol/L glucose and supplemented with 10 mmol/L nicotinamide, 10 mmol/L HEPES, 0.25 μg/mL Fungizone, 50 μg/mL gentamicin, 2 mmol/L glutamine, 10 g/mL ciprofloxacin, and 10% FCS. Following isolation and purification, both mouse and human islets were cultured for 1–4 days in RPMI 1640 medium containing 5.5 mmol/L glucose and supplemented with 10% FCS, 100 μg/mL penicillin, 100 μg/mL streptomycin, and 1% BSA. Palmitate (0.5 mmol/L) was present for 1 or 48 h prior to the microscopy recordings. A 100 mmol/L stock solution of palmitate was prepared in 50% ethanol. This solution was subsequently diluted in culture medium to a final concentration of 0.5 mmol/L and then allowed to complex with fatty acid–free BSA at 37°C. Control groups were exposed for the same period of time to media without palmitate, but otherwise with identical composition, including ethanol.
The islets were infected with adenovirus encoding GFP4-Grp1 or the cAMP biosensor using 105 fluorescent forming units/islet in culture medium as previously described (23). After 1 h incubation at 37°C, the inoculum was removed and the islets were washed and further cultured for 20–24 h. Before microscopy recordings, the islets were incubated for 30 min at 37°C in experimental buffer containing 125 mmol/L NaCl, 4.8 mmol/L KCl, 1.3 mmol/L CaCl2, 1.2 mmol/L MgCl2, and 25 mmol/L HEPES with pH adjusted to 7.40 with NaOH. They were then allowed to attach to the center of poly-lysine-coated round 25-mm glass coverslips for 5 min. β-Cells were identified based on their large size and negative response to adrenaline (23).
Rat insulinoma INS1E cells (26) were used between passages 65–90 and cultured in a humidified atmosphere containing 5% CO2 in RPMI 1640 medium containing 11 mmol/L glucose and supplemented with 10 mmol/L HEPES, 10% (v/v) heat-inactivated FBS, 2 mmol/L glutamine, 100 U/mL penicillin, 100 µg/mL streptomycin, 1 mmol/L sodium pyruvate, and 50 µmol/L 2-mercaptoethanol. For imaging experiments, cells were seeded onto poly-lysine-coated 25-mm coverslips, and transfections were performed with 0.2–0.5 µg of plasmid and 0.5–1 µg Lipofectamine 2000 in 100 µL Opti-MEM-I per coverslip. Cells were then further cultured in RPMI 1640 medium for 48 h in the absence or presence of 0.5 mmol/L palmitate in 1% (3:1) or 0.5% (6:1) of BSA.
Insulin-secreting MIN6 β-cells of passages 17–30 (27) were cultured in DMEM containing 25 mmol/L glucose and supplemented with 15% FCS, 2 mmol/L glutamine, 70 µmol/L 2-mercaptoethanol, 100 U/mL penicillin, and 100 µg/mL streptomycin. Where indicated, MIN6 cells were transfected with 0.5 µmol/L Lipofectamine 2000 and 100 nmol/L small interfering RNA (siRNA) against adenylyl cyclase (AC) 9 48 h prior to continued experimental handling.
Assessment of Cell Viability
Apoptosis in INS1E cells was assayed by the Cell Death Detection ELISAPLUS kit (Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s instructions. This photometric enzyme immunoassay measures cytoplasmic mono- and oligonucleosomes that increase after apoptosis-associated DNA degradation. The apoptosis measurements were related to the DNA content obtained in separate experiments and compared with the average value obtained from control cells.
The viability of cells within intact mouse islets was assessed by incubating islets for 15 min in experimental buffer supplemented with 10 µg/mL propidium iodide (Sigma) and 2 µmol/L calcein acetoxymethyl ester (Life Technologies) followed by imaging with a confocal microscope (described by Idevall-Hagren et al. ). The area of propidium iodide stained nuclei in relation to the calcein-stained viable cells was calculated with ImageJ (29).
Batch Measurements of Insulin Secretion
Control and palmitate-treated islets were preincubated in experimental buffer for 30 min at 37°C. Groups of 10 islets were then transferred to the different test media and incubated for 30 min. After collection of supernatants, the islets were sonicated for 10 s in acid ethanol to extract total insulin. The sample contents of insulin were determined by an electrochemiluminescence immunoassay (Meso Scale Diagnostics, Gaithersburg, MD).
RNA Isolation and Real-Time PCR
Total RNA was extracted from islets using the RNEasy micro kit (Qiagen, Hilden, Germany). Real-time PCR was performed using the Lightcycler instrument (Roche) and the Quanti Tect SYBR Green RT-PCR kit (Qiagen, Hilden, Germany) using the following primers: mouse AC1, forward 5′-ctctactaccagtcctactc-3′, reverse 5′-cttatagaagtccttgtccat-3′; human AC1, forward 5′-cagtacgacatctggggaaacac-3′, reverse 5′-agtcacctggattctgccctg-3′; mouse AC3, forward 5′-tgaggagagcatcaacaacg-3′, reverse 5′-tggtgtgactcctgaagctg-3′; human AC3, forward 5′-tgcacactcatggactttag-3′, reverse 5′-caggcacacacaggaatag-3′; mouse AC5, forward 5′-aacgactccacctatgacaa-3′, reverse 5′-aatgactccagccactacag-3′; human AC5, forward 5′-aggatgtgtttccctcatttc-3′, reverse 5′-ccatgcatcctcctacttaatc-3′; mouse AC6, forward 5′-tatgccgctatcttcctgct-3′, reverse 5′-tggcagagatgaacacaagc-3′; human AC6, forward 5′-ggccatcaggtacttcatta-3′, reverse 5′-caccaagtagtctcttcctatc-3′; mouse AC8, forward 5′-gtcaggaaggacaacacctc-3′, reverse 5′-tgtaggtggcgaagagtgta-3′; human AC8, forward 5′-agtactttgtcttcacggggg-3′, reverse 5′-gcaatcatgatcagcagcact-3′; mouse AC9, forward 5′-catacagaaggcaccgatag-3′, reverse 5′-ccgaacaggtcattgagtag-3′; human AC9, forward 5′-tgcggaatcacatttgctcc-3′, reverse 5′-gccttggtgctgtttttatgc-3′; mouse PDE8B, forward 5′-gactgatgaagagaagag-3′, reverse 5′-atgtctgttatgaagtagt-3′; PDE1C, forward 5′-aagcagcagaacggtgactt-3′, reverse 5′-ggcaaggtaatgcgacttgt-3′; PDE3B, forward 5′-ccaattcctggcttacctca-3′, reverse 5′-gtgatcgtaatcgtgcatgg-3′; PDE4A, forward 5′-catcaatgtcccacgatttg-3′, reverse 5′-taagtcccgctcctggaata-3′; mouse β-actin, forward 5′-gttacaggaagtccctcacc-3′, reverse 5′-ggagaccaaagccttcatac-3′; human β-actin, forward 5′-gggcatgggtcagaaggatt-3′, reverse 5′-tcgatggggtacttcagggt-3′; human GAPDH, forward 5′-aattccatggcaccgtcaag-3′, reverse 5′-gatctcgctcctggaagatgg-3′. Mouse PCR products were normalized to β-actin, and expression levels are given relative to control according to the following formula: fold change = 2−ΔΔCt, where ΔΔCt = (CtAC/PDE test − Ctβ-actin test) − (CtAC/PDE control − Ctβ-actin control). For human ACs, PCR efficiency was calculated from a standard curve generated from experiments on total RNA from EndoC-βH1-cells (generous gift from Nils Welsh, Uppsala University). The relative expression is given according to the following formula: fold change = (EAC)ΔCtAC / (Eactin)ΔCtβ-actin, where E = efficiency from standard curve; ΔCtAC = CtAC control – CtAC test; and ΔCtβ-actin = Ctβ-actin control – Ctβactin test.
Recordings of [cAMP]pm and Plasma Membrane PtdIns(3,4,5)P3
Measurements of fluorescence from the CFP/YFP-based [cAMP]pm reporter or the GFP-based PtdIns(3,4,5)P3 sensor were performed with total internal reflection fluorescence (TIRF) microscopy using either a custom-built prism-based system (22) or an objective-based setup (23). The prism-type TIRF setup was built around an E600FN upright microscope (Nikon Corp., Tokyo, Japan). A helium-cadmium laser (Kimmon, Tokyo, Japan) provided 442 nm light for excitation of CFP, and the 514 nm line of an argon laser (ALC 60×, Creative Laser Production, Munich, Germany) was used to excite YFP. Interference filters (Semrock, Rochester, NY) mounted in a filter wheel (Sutter Instruments, Novato, CA) were used to select the appropriate wavelength. The merged laser beam was homogenized and expanded by a rotating light-shaping diffuser (Physical Optics Corp., Torrance, CA) and refocused through a modified quartz dove prism (Axicon, Minsk, Belarus) with a 70° angle to achieve total internal reflection. The experimental chamber with cells was mounted on the custom-built stage of the microscope such that the cover slip was maintained in contact with the dove prism by a layer of immersion oil. Fluorescence light was collected through a 40×, 0.8-NA water immersion objective (Nikon). The objective-based system consisted of an Eclipse Ti microscope (Nikon) with a TIRF illuminator (Nikon) and a 60×, 1.45-NA objective. The 458-, 488-, and 514-nm lines of an argon laser (ALC60×, Creative Laser Production) were used to excite CFP, GFP, and YFP, respectively. The beam was coupled to the TIRF illuminator through an optical fiber (Oz Optics, Ottawa, Canada). In both evanescent wave microscope setups, fluorescence was detected with backilluminated electron-multiplying charge-coupled device cameras (iXON DU-897, Andor Technology, Belfast, Northern Ireland) under MetaFluor (Molecular Devices Corp., Downington, PA) software control. Emission wavelengths were selected with filters (485/25 nm half-bandwidth for CFP, 527/27 nm for GFP, and 560/40 nm for YFP [Semrock Rochester, NY]) mounted in a filter wheel (Sutter Instruments). For time-lapse recordings, images or image pairs were acquired every 5 s. To minimize exposure of the cells to the potentially harmful laser light, the beam was blocked by a mechanical shutter (Sutter Instruments) between image captures.
Recordings of the Cytoplasmic Ca2+ Concentration
For measurements of the cytoplasmic Ca2+ concentration ([Ca2+]i), cells and islets were preincubated during 30–40 min in the presence of 1 µmol/L of the acetoxymethyl esters of the Ca2+ indicators Fura-PE3 or Fluo-5F. Imaging of the Fura-PE3-loaded cells was performed with an inverted microscope (Diaphot, Nikon) equipped with a 40× 1.3-NA objective and an epifluorescence illuminator (Cairn Research Ltd., Faversham, U.K.) connected through a 5-mm diameter liquid light guide to an Optoscan monochromator (Cairn Research Ltd.) with a 150-W xenon arc lamp. The monochromator provided excitation light at 340 and 380 nm that was reflected by a 400-nm dichroic beam splitter, and emission was measured at 510/40 nm half-bandwidth using a Cascade (Photometrics, Tucson, AZ) or iXON DU-897 (Andor Technology) electron-multiplying charge-coupled device camera. The MetaFluor software (Molecular Devices) controlled the monochromator and the camera, acquiring image pairs every 2 s with 100–400 ms integration at each wavelength and <1 ms for changing wavelength and slits. To minimize bleaching and photodamage, the monochromator slits were closed until the start of the next acquisition cycle. Ratio images (340/380 nm) were obtained after subtraction of background and [Ca2+]i values calculated as previously described (30). Fluo-5F–loaded islets were imaged using the objective-based TIRF system described above for the PtdIns(3,4,5)P3 measurements.
Image analysis was made using MetaFluor (Molecular Devices). [cAMP]pm was expressed either as the ratio of CFP over YFP fluorescence after background subtraction with the basal ratio normalized to 1 to compensate for variability in expression levels, or as the Cα-YFP fluorescence intensity (F) in relation to the initial fluorescence intensity (F0) after subtraction of background. Subplasma-membrane [Ca2+]i and the membrane PtdIns(3,4,5)P3 concentration were evaluated as relative changes of the background-corrected Fluo-5F and GFP4-Grp1 fluorescence intensities, respectively (F/F0). Time-average levels of the second messengers were calculated by measuring the area under curve followed by normalization for the elapsed time. Data are presented as mean ± SEM. The primary cell data were obtained from at least three independent islet preparations. The apoptosis data were statistically analyzed with one-way ANOVA followed by Tukey multiple comparison test, and the analyses of fraction-responsive cells were made with the χ2 test. In all other cases statistical significances were evaluated using Student t test.
Long-term Exposure to Palmitate Suppresses Glucose-Induced cAMP Signaling in β-Cells
[cAMP]pm was monitored in INS1E β-cells as well as in superficially located β-cells within intact mouse and human islets. As previously reported for MIN6 and primary mouse β-cells (21,23), INS1E cells responded to an increase in the glucose concentration with pronounced [cAMP]pm oscillations (Fig. 1A). Subsequent addition of 100 nmol/L GLP-1 caused stable [cAMP]pm elevation. After 48-h exposure to 0.5 mmol/L palmitate in 0.5 or 1% BSA (fatty acid:BSA molar ratio of 6:1 and 3:1, respectively), the glucose-induced [cAMP]pm response was significantly delayed and the amplitude suppressed, whereas the amplitude and duration of GLP-1–triggered [cAMP]pm elevation was unaffected (Fig. 1B–F). The palmitate treatments caused modest but significant (P < 0.01 at 3:1; P < 0.001 at 6:1) increases of INS1E cell apoptosis (Fig. 1G).
Similarly, mouse β-cells in intact islets responded to an increase of the glucose concentration from 3 to 20 mmol/L with shortly delayed and pronounced [cAMP]pm elevation, often with oscillations with frequencies in the 0.09–0.27/min range (Fig. 2A). Addition of 100 nmol/L GLP-1 triggered stable [cAMP]pm elevation, which was counteracted by 5 µmol/L adrenaline used to confirm the β-cell identity (23) (Fig. 2A). After 48-h exposure to 0.5 mmol/L palmitate in 1% BSA, the glucose-induced [cAMP]pm elevation, but not that triggered by GLP-1, was significantly suppressed (Fig. 2B and E) (P < 0.001). GLP-1 was also tested at 100 pmol/L, and also at this concentration, there was no difference in [cAMP]pm between palmitate-treated islets and control (time-average CFP/YFP 1.79 ± 0.35 vs. 1.98 ± 0.22; n = 35 and 13 cells, respectively, from five islets in three independent experiments). The poor glucose response was not due to initial cAMP elevation by the palmitate treatment since there was no cAMP-lowering effect of the AC inhibitor 2′,5′-dideoxyadenosine (100 µmol/L; data not shown). The impaired glucose response was not due to reduced islet viability. Accordingly, the ratio of propidium iodide to calcein-labeled cell areas were similar in control (0.0027 ± 0.0005; n = 36 islets) and palmitate-treated islets (0.0036 ± 0.0011; n = 30 islets) and hundred-fold lower than in heat-shocked control islets incubated for 5 min at 50°C (0.2668 ± 0.0297; n = 50 islets).
Human islet β-cells behaved essentially as mouse β-cells. At 3 mmol/L glucose, [cAMP]pm was low and stable in 57% of the cells (n = 14), and in the remaining cells, there were modest [cAMP]pm fluctuations (not shown). When the glucose concentration was increased to 20 mmol/L, [cAMP]pm increased and oscillated (frequency 0.08–0.24/min; amplitude 0.16 ± 0.03 ratio units; n = 37 oscillations from 8 cells) often from an elevated level (Fig. 2C). GLP-1 (100 nmol/L) induced sustained [cAMP]pm elevation, which was reversed by 5 µmol/L adrenaline (Fig. 2C). Like in INS1E and mouse β-cells, 48-h exposure to palmitate selectively suppressed the glucose-induced [cAMP]pm elevation (P < 0.001) (Fig. 2D and E).
Short Palmitate Treatment Does Not Affect Glucose- and GLP-1–Induced [cAMP]pm Signaling
Acute exposure of mouse islet β-cells to 0.5 mmol/L palmitate (1% BSA) in the presence of 3 mmol/L glucose sometimes resulted in transient [cAMP]pm elevation (7 out of 15 cells) (Fig. 3A). However, there was no effect when palmitate was added in the presence of 20 mmol/L glucose or 100 nmol/L GLP-1 (Fig. 3B-C). Likewise, there were no differences in glucose- or GLP-1–induced cAMP signaling after 1-h preincubation with palmitate (Fig. 3D–F). The mechanism behind the modest [cAMP]pm elevation triggered by acute palmitate treatment is not known but may be secondary to the elevation of [Ca2+]i by fatty acid–mediated activation of GPR40 receptors (31).
Palmitate Does Not Impair Glucose-Induced [Ca2+]i Signaling
Since glucose-induced cAMP signaling is amplified by Ca2+ (23), we next investigated whether [Ca2+]i is affected by long-term exposure to palmitate. In both INS1E and mouse islet β-cells loaded with Fura-PE3, the basal [Ca2+]i at substimulatory glucose concentrations was slightly, but significantly (P < 0.05), increased after exposure to palmitate, but the fatty acid influenced neither the ability of 20 mmol/L glucose to induce [Ca2+]i oscillations, the [Ca2+]i elevation induced by K+ depolarization (Fig. 4A–D, G, and H), nor the synchronization of oscillations among neighboring β-cells (not shown). TIRF recordings of subplasma-membrane [Ca2+]i with the low-affinity indicator Fluo-5F also did not reveal any influence of palmitate on [Ca2+]i (Fig. 4E, F, and I).
Palmitate Impairs Glucose-Induced Pulsatile Insulin Secretion
To investigate if the compromised [cAMP]pm signaling was associated with impaired insulin secretion, we monitored the time-course of insulin secretion from individual cells within intact islets by detecting the lipid PtdIns(3,4,5)P3, which is formed in the plasma membrane following autocrine insulin receptor activation (21). Control mouse and human islet β-cells responded to glucose with pronounced PtdIns(3,4,5)P3 oscillations, reflecting pulsatile insulin secretion (Fig. 5A and B). The glucose response was almost abolished after 48-h palmitate treatment, and the cells showed only a transient increase or minute stable PtdIns(3,4,5)P3 elevation without oscillations (Fig. 5C–E). The reduced response was not due to impaired insulin signaling since exogenously added insulin evoked PtdIns(3,4,5)P3 elevation (Fig. 5C–E). Consistent with deficient cAMP production underlying the impaired secretory response to glucose, the fraction of cells responding with increased PtdIns(3,4,5)P3 was doubled and the response amplitude several-fold increased when 20 mmol/L glucose was added in the presence of the AC activator forskolin (Fig. 5E and F). The palmitate-induced suppression of glucose-induced insulin secretion was confirmed by measurements of insulin release from batch-incubated islets using a conventional immunoassay (Fig. 5G). The results also demonstrate that 1 nmol/L GLP-1 restores insulin secretion to similar levels as in the control (Fig. 5G).
Reduction of AC Expression Underlies Palmitate Impairment of Glucose-Induced cAMP Production
Next, the expression of ACs and phosphodiesterases (PDEs), synthesizing and degrading cAMP, respectively, was evaluated by real-time PCR. After 48-h palmitate treatment of mouse islets, AC9 mRNA was reduced to 70% (P < 0.01), but there were no changes in expression of AC1, AC3, AC5, AC6, or AC8 (Fig. 6A) or any of the tested PDEs (PDE1C, −3B, −4A, and −8B) (Fig. 6B). In human islets, AC5 was reduced and AC8 was increased, while other ACs did not show significant alterations in mRNA expression (Fig. 6C). The importance of AC9 for glucose-induced [cAMP]pm signaling in mouse β-cells was investigated by siRNA-mediated knockdown in MIN6 cells. After 48-h treatment with siRNA, AC9 mRNA was reduced to 50%, and only 21% of the cells responded to glucose with [cAMP]pm oscillations as compared with 91% in control-siRNA-treated cells (Fig. 6D and E). The average glucose-induced [cAMP]pm response was thus significantly suppressed after AC9 knockdown (Fig. 6F) (P < 0.001). Also the GLP-1–induced [cAMP]pm elevation was reduced, but not to the same extent (Fig. 6F) (P < 0.05). The reduced AC9 expression was accompanied by impaired insulin secretion. When evaluating glucose-induced PtdIns(3,4,5)P3 formation, the initial 10 min of the glucose response was unaffected, whereas the subsequent sustained response was significantly reduced in siRNA-treated cells (Fig. 6G, H, and K). The lower insulin secretion most likely reflected the deficient cAMP production since the response was restored by addition of 1 mmol/L 8-Br-cAMP (Fig. 6I–K).
Deleterious effects of free fatty acids on β-cell function are well documented (5–8), but the mechanisms underlying the defective insulin secretion are not well understood. In the current study, we show that inhibition of glucose-induced cAMP production in β-cells contributes to impaired insulin secretion after long-term exposure to palmitate.
With the advent of techniques to monitor cAMP dynamics in single cells, increasing evidence indicates that intracellular cAMP elevation is an integral part of glucose stimulus-secretion coupling in β-cells (21,32,33). Measurements in the subplasma-membrane space of both clonal and primary mouse β-cells have shown that cAMP often oscillates and contributes to pulsatile release of insulin (21–23). The present data show that glucose triggers cAMP oscillations also in human β-cells. Increases of cAMP amplify insulin secretion both via PKA and the guanine nucleotide exchange factor Epac. The cAMP effectors act at multiple levels to influence the activity of various ion channels and other proteins involved in signaling and exocytosis (34–40). It is therefore not surprising that deteriorated glucose-induced cAMP generation results in impaired insulin secretion.
The observation that glucose-induced cAMP signaling is suppressed in palmitate-treated cells is consistent with previous findings that the secretory defect involves a late step in stimulus-secretion coupling (16). The poor cAMP response was not secondary to changes in [Ca2+]i, which was essentially unaffected by palmitate exposure, except for slight elevation of basal [Ca2+]i also observed in previous studies (16). The upstream events with metabolic generation of ATP, membrane depolarization, and Ca2+ influx therefore seem unaffected, which is consistent with palmitate lacking effect on glycolytic flux and ATP synthesis in MIN6 and mouse islet β-cells (16,41).
It has been suggested that palmitate impairs insulin secretion via PKA-mediated induction of the cAMP early repressor (ICER-1γ) and suppression of connexin-36 expression (42), and lipotoxicity has indeed been associated with impaired coordination of Ca2+ signals in human islets (43). The increased PKA activity suggested to underlie the altered connexin-36 expression is difficult to reconcile with the present observation of reduced cAMP production. Moreover, we did not observe any striking deterioration of the synchronization of glucose-induced [Ca2+]i oscillations among neighboring β-cells. The impaired insulin secretion after chronic palmitate exposure has also been attributed to dissociation of the secretory vesicles from the Ca2+ entry sites (17). Our data are more easily reconciled with such a mechanism. Accordingly, the protein complex responsible for tethering of secretory vesicles to Ca2+ channels includes components that are directly regulated by cAMP. For example, the SNARE proteins syntaxin and SNAP25 are both regulated by PKA, and Epac interacts with the Rab-binding protein RIM2, which, in turn, interacts with the L-type voltage-dependent Ca2+ channel (37,44,45). Defective cAMP generation might thus affect the interaction between Ca2+ channels and the secretory granules.
Reduced expression of ACs provides a plausible explanation for the defective glucose-induced cAMP responses. In human islets, AC5 was reduced by palmitate treatment. Interestingly, this enzyme was recently demonstrated to be important for glucose-induced insulin secretion in human β-cells (46). In the mouse, AC9 seems to be more important. Although the activity of this AC typically is low (47), its functional importance was verified in siRNA knockdown experiments in MIN6 cells. Further studies are warranted to clarify the detailed involvement in insulin secretion of the many ACs expressed in mouse and human islets. Moreover, it cannot be excluded that additional factors contribute to the palmitate-induced impairment of cAMP signaling, such as altered expression of additional cAMP-regulating proteins or direct modulation of protein function via, e.g., acylation or palmitoylation (48,49).
β-Cell dysfunction induced by long-term exposure to palmitate typically culminates in apoptotic cell death (7,12). However, our relatively mild treatment regimen had little effect on cell viability, indicating that the observed signaling defect represents an early event in the lipotoxic process causing β-cell failure. Importantly, the deficient glucose-induced cAMP production did not reflect a general deterioration of the cAMP signaling system since the cells responded normally to GLP-1. cAMP has been found to protect from apoptosis in many cell types, including palmitate-induced apoptosis in insulin-secreting cells (50). Impaired glucose-induced cAMP generation might therefore not only explain defective insulin secretion but also contribute to the general β-cell dysfunction after chronic exposure to palmitate.
Acknowledgments. The authors thank Claes Wollheim and Pierre Maechler, University of Geneva, for the gift of INS1E cells; Junichi Miyazaki, Kobe University, for providing MIN6 cells; and Parvin Ahooghalandari, Uppsala University, for technical assistance.
Funding. This work was supported by grants from the Swedish Research Council, the European Foundation for the Study of Diabetes/Merck Sharp & Dohme, the Novo Nordisk Foundation, the Swedish Diabetes Foundation, the Family Ernfors Foundation, the Swedish national strategic grant initiative EXODIAB (Excellence of Diabetes Research in Sweden), the National Natural Science Foundation of China (81400771), and Taishan Scholars Construction Engineering and the Science and Technology Project for the Universities of Shandong Province (J14LE01). Human pancreatic islets were obtained from the Nordic Network for Clinical Islet Transplantation, supported by EXODIAB and JDRF.
Duality of Interest. No potential conflicts of interest relevant to this article were reported.
Author Contributions. G.T. researched and analyzed data and wrote the manuscript. E.M.S., Y.X., and H.S. researched and analyzed data. A.T. designed the study, analyzed data, and wrote the manuscript. A.T. 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.
Prior Presentation. Parts of this study were presented at the 49th Annual Meeting of the European Association for the Study of Diabetes, Barcelona, Spain, 23–27 September 2013.