Paracrine control of α-cell glucagon exocytosis is compromised in human type-2 diabetes

Glucagon is released from pancreatic α-cells to activate pathways that raise blood glucose. Its secretion is regulated by α-cell-intrinsic glucose sensing and paracrine control through insulin and somatostatin. To understand the inadequately high glucagon levels that contribute to hyperglycemia in type-2 diabetes (T2D), we analyzed granule behavior, exocytosis and membrane excitability in α-cells of 68 non-diabetic and 21 T2D human donors. We report that exocytosis is moderately reduced in α-cells of T2D donors, without changes in voltage-dependent ion currents or granule trafficking. Dispersed α-cells have a non-physiological V-shaped dose response to glucose, with maximal exocytosis at hyperglycemia. Within intact islets, hyperglycemia instead inhibits α-cell exocytosis, but not in T2D or when paracrine inhibition by insulin or somatostatin is blocked. Surface expression of somatostatin-receptor-2 is reduced in T2D, suggesting a mechanism for the observed somatostatin resistance. Thus, elevated glucagon in human T2D may reflect α-cell insensitivity to paracrine inhibition at hyperglycemia. Glucagon is elevated Type-2 diabetes, which contributes to poor glucose control in patients with the disease. Here the authors report that secretion of the hormone is controlled by paracrine inhibition, and that resistance of α-cells to somatostatin can explain hyperglucagonemia in type-2 diabetes.

G lucagon is released from pancreatic α-cells and counteracts the glucose-lowering actions of insulin by stimulating gluconeogenesis and hepatic glucose output. Initially thought of only as part of the body's defense against hypoglycemia 1 , it is now clear that inadequate glucagon levels also contribute to diabetic hyperglycemia and present a challenge for diabetes management 2,3 . Glucagon secretion is triggered by low blood glucose and suppressed at physiological glucose levels, and both α-cell intrinsic and paracrine mechanisms have been cited to explain these effects. In the intrinsic models, glucose metabolism and the generation of ATP play a central role [4][5][6] , either through subtle, K ATP channel-dependent depolarization of the resting membrane potential and subsequent inactivation of Na + -channels 7-10 , or as consequence of glucose-induced activation of the sarco/endoplasmic reticulum Ca 2+ -ATPase that leads to closure of store-operated channels and hyperpolarization 11,12 . In addition, intrinsic glucose-dependent cAMP signaling may play a role 13 . However, none of these models fully explain the glucose concentration dependence of glucagon secretion, in particular in the hyperglycemic range.
Glucagon secretion is also under paracrine control from neighboring βand δ-cells, and the inhibitory effects of somatostatin [14][15][16][17][18] , insulin [19][20][21][22][23] , and GABA 24,25 have long been recognized. Paracrine inhibition is likely to play a role at elevated glucose levels, when βand δ-cells are active. Indeed, glucagon is secreted in pulses that are anti-synchronous to pulses of insulin and somatostatin 26,27 . This relationship is important for the postprandial suppression of glucagon secretion and is lost in type-2 diabetes and pre-diabetes [28][29][30] . α-cells express the somatostatin receptor SSTR2, which leads to hyperpolarization via activation of GIRK-channels 31,32 . Somatostatin also inhibits the exocytosis machinery via calcineurin 32 , and inhibits α-cell exocytosis by effectively decreasing cytosolic cAMP 33,34 . Insulin receptor signaling is required for the suppression of glucagon secretion in vivo 35 , but the precise mechanisms behind this are still debated 20,34,36 . Decreased sensitivity to insulin (or somatostatin) may therefore underlie the inadequate glucagon secretion in type-2 diabetes 37 .
Glucagon is stored in~7000 granules (diameter~270 nm) and secreted by Ca 2+ -and SNARE protein-dependent exocytosis 38,39 . At any time, only~1% of these granules are in a releasable state that can undergo exocytosis upon Ca 2+ -influx 40 . Paracrine signaling and glucose regulate glucagon secretion at least in part by affecting the size of this releasable pool of granules 32,38,41 . In many endocrine cells, secretory granules become release ready by sequential docking at the plasma membrane and assembly of the secretory machinery (priming) 42,43 . Although disturbances in these steps have been documented in β-cells of type-2 diabetic donors 44-47 , they have not yet been studied in α-cells. An obstacle for understanding the regulation of α-cells has been the difficulty to isolate intrinsic and paracrine factors of α-cell regulation, as well as species differences between humans and rodent models. Glucagon secretion in vivo and in intact islets is affected by the presence of neighboring cell types, while single-cell electrophysiological measurements are invasive and may not reflect the in vivo situation.
In the current work, we took an optical approach to study glucagon granule exocytosis in α-cells of non-diabetic (ND) and type-2 diabetic (T2D) human subjects. We report that α-cells within intact islets respond with physiological inhibition of exocytosis by elevated glucose, whereas dispersed α-cells have a V-shaped response to glucose due to the lack of paracrine inhibition by insulin and somatostatin from neighboring βand δ-cells. Importantly, α-cells of T2D are resistant to inhibition by insulin and somatostatin, which might underlie the hyperglucagonemia in type-2 diabetes.

Results
Exocytosis of glucagon granules in human α-cells. Docking and exocytosis of glucagon granules at the plasma membrane was studied in dispersed islet preparations from 68 non-diabetic (ND) donors that all had glycated hemoglobin HbA1c values <6% (average 5.57 ± 0.29%, Supplementary Fig. 5). To identify α-cells, we transduced with Pppg-EGFP and the secretory granule marker NPY-mCherry, or with Pppg-NPY-EGFP ( Fig. 1a and S1a, b) to drive expression of fluorescent proteins from the pre-proglucagon promoter (see methods). After culture for 26-48 h, α-cells were imaged by total internal reflection (TIRF) microscopy, which selectively images fluorescence near the plasma membrane (exponential decay constant τ~0.1 µm). The granule marker had a punctate staining pattern and excellent overlap with antiglucagon immunostaining (Fig. 1a). Local application of elevated K + (75 mM, replacing Na + ) to depolarize the cells resulted in exocytosis, seen as rapid disappearance of individual fluorescently labeled granules (gr, see Fig. 1b and examples in S1B,C). Exposure to elevated K + for 40 s released 0.078 ± 0.004 granules µm −12 (169 α-cells/29 ND donors, Fig. 1c, black). Exocytosis proceeded initially with a burst (5.2 × 10 −3 gr µm −2 s −1 during the first 10 s) and decreased later to <0.6 × 10 −3 gr µm −2 s −1 ; these rates are about one-third of those observed in human β-cells 44 . Fitting the cumulative exocytosis (n = 1530 granules) with a double exponential function revealed two components with time constants of τ = 3.6 ± 0.2 s and 19.9 ± 0.9 s (Fig. 1c). The faster component made up 39 ± 3% of the total response, and likely corresponds to the RRP. Exocytosis occurred in granules that from the start of the experiment had been docked at the plasma membrane. We therefore quantified changes in docked granules during the experiment, by measuring the density of granules in the TIRF field. Stimulated exocytosis partially depleted docked granules (Fig. 1d), indicating that replacement by docking of new granules is relatively slow (a notion confirmed by a double stimulation protocol, Supplementary Fig. 2). On average, 13 ± 0.7% of the docked granules were released during the stimulation. Thus, depolarization of α-cells results in exocytosis with biphasic kinetics similar to those in other endocrine cells, indicating the existence of granule pools with differing release probabilities.
Voltage-dependent currents are normal in T2D α-cells. Since exocytosis in α-cells depends on Ca 2+ -influx, we characterized voltage-dependent ion currents using patch-clamp electrophysiology. Dispersed ND or T2D α-cells were voltage clamped in whole-cell mode, and subjected to step depolarizations up to +70 mV from a holding potential of −70 mV (Fig. 2a). Analysis of the resulting inward currents revealed peak Ca 2+ (Fig. 2b) and Na + -currents (Fig. 2c) that were of similar amplitude in ND and T2D cells. Half-maximal Ca 2+ -current activation was reached at −23 ± 0.4 (ND) and −25 ± 1.4 mV (T2D, n.s.), and half-maximal Na + -current activation was at −25 ± 0.6 (ND) and −24 ± 0.8 mV (T2D, n.s.). We also determined depolarization evoked membrane capacitance increases, a measure of exocytosis (Fig. 2d, e). A train of 14 depolarizations to 0 mV lasting 200 ms each resulted in a total capacitance increase of 112 ± 19 fF in ND cells and 78 ± 19 fF in T2D cells. This corresponds to a reduction of exocytosis by 25 ± 10% in T2D (p = 0.1), which is similar to the reduction observed by imaging granule release (Fig. 1d). Cell size, as assessed by cell capacitance was not different in the two groups (Fig. 2f). Thus, reduced exocytosis in T2D α-cells cannot be explained by changes in Ca 2+ -channel behavior.
Glucose regulation of glucagon secretion. Next, we determined the physiological glucose dependence of dispersed α-cells by measuring spontaneous exocytosis in a range of ambient glucose concentrations (1, 3, 7, 10, or 20 mM; at least 20 min pre-incubation), without imposing any depolarization. In movies lasting 3 min, we quantified granule exocytosis, docked granules, and the rate of docking of new granules at the plasma membrane ( Fig. 3a-d). Spontaneous exocytosis was observed in all glucose concentrations, with a bimodal (V-shaped) dose response to glucose. Inhibition was about half at the nadir of 7 mM glucose, and ND and T2D cells behaved essentially identically (Fig. 3b). Docked granules, and to a lesser degree the rate of docking (in the same cells) likewise had a bimodal response to glucose, with a nadir at 7 mM (Fig. 3a, c, d). In separate experiments, we noticed that the response to changes in the glucose concentration was slow, and only minor during 2 min observation ( Supplementary Fig. 3a, b). Thus, the glucose dependence of dispersed α-cells does not reflect physiological glucagon secretion, and exocytosis is accelerated in the hyperglycemic range instead of being inhibited. The data also suggest that the availability of docked granules is part of the regulation of exocytosis in α-cells.
To experimentally isolate direct glucose effects on the exocytosis machinery, we applied K + -stimulations to α-cells bathed in 1, 7, or 10 mM glucose (Fig. 3e). This approach primarily elicits exocytosis of granules that are "primed" for exocytosis, the readily releasable pool (RRP). At all glucose concentrations, elevated K + stimulated biphasic exocytosis that by far exceeded spontaneous exocytosis (Fig. 3e). However, we observed again a bimodal glucose dependence with a nadir at 7 mM (38 cells/6 donors), where K + -induced exocytosis was reduced by about half compared with 1 mM glucose (−45 ± 9%, p = 8 × 10 −4 , 30 cells/6 donors), or 10 mM glucose (−47 ± 6%, p = 5 × 10 −6 , 71 cells/14 donors). This reduction was most prominent during the initial burst phase, which may correspond to the immediately releasable pool of granules in β-cells 45 . At all glucose concentrations, K + -stimulated exocytosis was significantly slower in T2D α-cells than in ND, but the bimodal glucose dependence of docking and exocytosis was preserved (Fig. 3f). We conclude that the α-cell exocytosis machinery is regulated by the ambient glucose concentration.
Exocytosis in α-cells within intact islets. The unexpected Vshaped glucose response of dispersed α-cells indicates that intrinsic regulation cannot explain the physiological inhibition of glucagon secretion in hyperglycemia. An alternative are paracrine effects from neighboring βand δ-cells, which prompted us to quantify glucose dependent exocytosis in α-cells within intact islets ( Fig. 4a, b). As expected, exocytosis of α-cells in intact islets varied with the glucose concentration, with inhibition in 7 mM glucose by 67 ± 15% (p = 0.016; 10 islets/3 donors), compared with 1 mM glucose (11 islets/3 donors). Exocytosis was also inhibited in 10 mM (by 56 ± 16% vs 1 mM, p = 0.024, 16 cells/4 donors), in contrast to dispersed α-cells. However, this inhibition was prevented by the SSTR2-specific somatostatin receptor antagonist CYN154806 (200 nM in the bath solution; p = 0.003 vs 10 mM glucose, 19 cells/3 donors). Similarly, block of insulin action with the insulin receptor antagonist S961 (1 μM) prevented inhibition of exocytosis at 10 mM glucose (p = 0.003 vs 10 mM glucose, 11 cells/2 donors), indicating that paracrine signaling is required for proper glucagon control in hyperglycemic conditions. In intact islets of T2D donors, α-cell exocytosis had a bimodal dose response to glucose that lacked inhibition hyperglycemia ( Fig. 4c). At 7 mM glucose (16 islets/4 donors), exocytosis was inhibited to about half compared with 1 mM (21 islets/5 donors), whereas 10 mM glucose has no effect (29 islets/5 donors). Thus, α-cells within intact ND islets have a physiological response to glucose. Disruption of paracrine signaling by antagonists or islet dispersion leads to a V-shaped glucose response, similar to that observed in intact T2D islets.
We confirmed 48 expression of SSTR2 in human α-cells by coimmunostaining the receptor and glucagon in pancreatic sections of 10 human donors (5 ND, 5 T2D; Fig. 4d). In ND islets, SSTR2 distribution was mostly confined to the cell membrane of αand other islet cells, whereas in T2D islets the SSTR2 staining was both weaker and largely vesicular (Fig. 4d). Quantitative analysis confirmed this conclusion and estimated that SSTR2 surface expression is decreased by 44 ± 7%, in T2D (824 cells/5 T2D donors vs 828 cells/5 ND donors Fig. 4e). Glucagon levels and distribution were similar in both groups (Fig. 4f). Insensitivity to somatostatin is therefore the result of excessive receptor internalization, as has recently been shown for pituitary cells 49 .

Rapid paracrine inhibition by insulin and somatostatin.
Glucagon secretion oscillates with a frequency of minutes, which is inconsistent with the relatively slow glucose dependent regulation ( Supplementary Fig 3). We therefore determined the time course of paracrine inhibition of spontaneous exocytosis by rapidly applying somatostatin (Fig. 6a, b, blue shading) or insulin (Fig. 6c, d, green shading) to dispersed α-cells. The effect frequently wore off towards the end of the 1 min challenge, and was rapidly reversed by removing the hormone (Fig. 6a). Similarly, rapid effects on exocytosis were observed with insulin (21 cells, 4 donors, Fig. 6c) or adrenaline ( Supplementary  Fig. 4). Thus, paracrine regulation of glucagon exocytosis occurs on a timescale that is consistent with the observed glucagon pulsatility 26 . Due to the small effect size, the time course of somatostatin or insulin inhibition of exocytosis could not be determined for T2D α-cells (Fig. 6b, d). Finally, we tested the effect of somatostatin and insulin on α-cell electrical activity by perforated patch-clamp electrophysiology (Fig. 6e, h). In 10 mM glucose, electrical activity consisted action potential trains as described previously 38,51 . In ND α-cells, pulses of somatostatin (400 nM) caused rapid cessation of electrical activity (Fig. 6e). In the example in Fig. 6e top, action potentials reappeared 20 s after removal of the somatostatin. However, in many cells electrical activity reappeared already during the somatostatin pulse, which is apparent the averaged membrane potential (Fig. 6e,  bottom). Similar recovery during the somatostatin pulse was also seen for exocytosis (Fig. 6a), and may reflect somatostatin receptor inactivation. Insulin also dampened electrical activity, but these effects were both slower and weaker than those of somatostatin (Fig. 6f). In α-cells from T2D donors, the somatostatin or insulin pulses had little effect on the time course of exocytosis (Fig. 6b, d) or electrical activity (Fig. 6g, h). In summary, insulin and somatostatin inhibit both exocytosis (Fig. 5) and electrical activity ( Fig. 6). Both effects are lost in T2D, which is consistent with the notion that these cells are resistant to paracrine inhibition.

Discussion
Glucose controls glucagon secretion by intrinsic and paracrine mechanisms, but their relative significance is still debated 52 , and secretory defects in type-2 diabetes are not well understood. The current work is first in using high-resolution microscopy to study glucagon secretion both in intact islets and in single dispersed α-cells of healthy and type-2 diabetic donors, thus isolating intrinsic from paracrine mechanisms while having full control over paracrine signaling. We show that in the absence of paracrine influence, isolated α-cells respond appropriately to hypoglycemia with an increase in glucagon granule exocytosis. This is consistent with the glucose dependence of glucagon secretion from intact islets (but not FACS sorted α-cells) 52,53 , and indicates that glucagon secretion in the lower glucose-concentration range is mostly under intrinsic control. With only 2-fold difference in the exocytosis rate between minimal secretion at 7 mM and maximal secretion at 1 mM glucose, the dynamic range is small compared with β-cells. Surprisingly, exocytosis of dispersed α-cells is stimulated in the hyperglycemic range, leading to an unphysiological V-shaped response with maximal exocytosis above 10 mM glucose. This is in contrast to intact islets, in which glucagon secretion is depressed between 3 and 20 mM glucose 54 . We confirm this here by exocytosis measurements in intact islets, and provide evidence that this depression depends on the inhibitory effects on insulin and somatostatin that are released by neighboring βand δ-cells. Evidently, α-cell intrinsic mechanisms are sufficient for the regulation of glucagon secretion hypo-and normoglycemic range (0-7 mM glucose), while paracrine inhibition is responsible for the physiological response in the hyperglycemic range. Consequently, appropriate glucagon secretion in the hyperglycemic range is lost when α-cells are removed from their context within the islet. Elevated glucagon is a hallmark of type-2 diabetes 55 . Despite this, both glucose-dependent and depolarization (K + )-induced exocytosis was reduced in α-cells from donors that had been diagnosed with T2D. Both exocytosis and docked granules were moderately anti-correlated with donor HbA1c values. This indicates that the exocytosis machinery in α-cells from type-2 diabetics is slightly impaired, while electrophysiological parameters (that determine electrical activity, depolarization and Ca 2+ -influx) were normal. The reason for this is unknown, but may reflect reduced expression of certain exocytosis-related proteins, as is the case in β-cells 44,46,47 . The reduced exocytotic capacity in T2D α-cells is unrelated to changes in electrical activity, because it could be observed in K + -stimulation experiments in which the membrane potential is clamped. Since exocytosis in single α-cells is impaired rather than increased in T2D, the hyperglucagonemia in diabetic humans must be due to mechanisms that are lost in isolated cells, such as paracrine or neuronal regulation 56 . In addition, gut derived glucagon may contribute to hyperglucagonemia following oral glucose intake 57,58 .
Strikingly, the inhibitory effects of insulin and somatostatin on glucagon exocytosis were strongly reduced in cells from T2D donors, in parallel with internalization and reduced surface expression of SSTR2, the major somatostatin receptor in human α-cells. This points to α-cell resistance to insulin and somatostatin as the main cause for inadequate glucagon secretion in type-2 diabetes, which in turn exacerbates hyperglycemia 59 . Insulin resistance is a hallmark of T2DM, and has previously been proposed as mechanism for hyperglucagonemia. For example, insulin resistance is associated with fasting glucagon levels 37 , and this inverse relationship is lost in type-2 diabetes 28 . Interestingly, SSTR2 surface expression was also reduced in β-cells within T2D islets, suggesting that reduced somatostatin sensitivity may contribute also to increased insulin secretion, as observed early during the development of T2D. While there is reason to believe that this is a consequence of altered δ-cells activity 60 , its role may be to adapt islets to periods of greater food availability.
Exposure to insulin, somatostatin, and GABA reduced α-cell exocytosis, while adrenalin and glutamate stimulated it. This is consistent with the known effects of these signaling molecules on islets, as well as systemically 52 . We show here that these effects are a b d c very rapid (seconds), which is consistent with the frequency of pulsatile glucagon release in vivo and from intact islets. In the absence of glucose-dependent control, paracrine inhibition by insulin and somatostatin is therefore the most likely mechanism for glucagon regulation in the hyperglycemic range. It can be speculated that the differential glucose dependence of insulin and somatostatin secretion is reflected in different target glucose ranges for their action on α-cells. All tested paracrine modulators affected exocytosis machinery at the priming step, rather than by increasing granule docking. This is consistent with previous findings that somatostatin inhibits exocytosis in rat α-cells through G i -dependent depriming 32 , and reports that antagonists of SSTR2 12 or the associated G-protein cascade 12 increase glucagon secretion without altering the glucose-dependent inhibition of glucagon secretion. We did not observe any bursts of exocytosis, as might be expected given the pulsatile glucagon secretion from intact islets. This is in line with the absence of membrane potential oscillations in single cells 61 (that we confirm here), and indicates that the islet context is required not just for intra-islet synchronization, but for oscillatory α-cell behavior as such.
Capacitance measurements indicate that glucagon granules exist in at least two states with different release probabilities, which are often referred to as the readily releasable pool of granules (RRP) and a larger reserve pool (RP) 38,62 . We show here that glucagon granules were present at the plasma membrane for extended periods before undergoing exocytosis. We interpret this as the relatively slow conversion from RP to RRP that reflects the molecular assembly of the secretory machinery at the release site, in analogy with the situation in β-cells 42,44 . Throughout the glucose range, the rate of exocytosis was nearly identical to that of granule docking (Fig. 1b-d), suggesting that docking is rate limiting for secretion. This may indeed be the case during strong (non-physiological) stimulation, as illustrated by the finding that the glucosedependence of depolarization-induced exocytosis followed that of granule docking. However, in physiological conditions elevated K + accelerated exocytosis~50-fold (during the first second), which indicates a large excess in exocytotic capacity that is not triggered by normal α-cell electrical activity. A possible explanation could be that only a limited number of granules is positionally primed, i.e. located near voltage-gated Ca 2+ -granules 63  work is required to understand the combination of factors affecting granule exocytosis, and the granule conversion rates provided here may be useful in this regard.

Methods
Tissue. Pancreatic islets and pancreas sections were obtained from human cadaveric donors by the Nordic Network for Clinical Islet Transplantation Uppsala 64 (ethical approval by Uppsala Regional Ethics Board) or the ADI Isletcore at the University of Alberta 65 (ethical approval by Alberta Human Research Ethics Board, Pro00001754), with written donor and family consent for use in research. Work with human tissue complied with all relevant ethical regulations for use of human tissue in research and the study was approved by the Uppsala Regional Ethics Board (2006/348). Isolated islets were cultured free-floating in sterile dishes in CMRL 1066 culture medium containing 5.5 mM glucose, 10% fetal calf serum (FCS), 2 mM L-glutamine, streptomycin (100 U/ml), and penicillin (100 U/ml) at 37°C in an atmosphere of 5% CO 2 up to 2 weeks. Islets were dispersed into single cells by gentle pipetting in cell dissociation buffer (Thermo Fisher Scientific) supplemented with trypsin (0.005%, Life Technologies). Cells were then washed and plated in serum-containing medium onto 22-mm polylysine-coated coverslips, allowed to settle overnight, and then transduced using adenovirus. In Fig. 4a, c, the dispersion step was omitted and intact islets were transduced with Pppg-NPY-EGFP adenovirus and allowed to settle onto 22-mm polylysine-coated coverslips.
Labeling of human pancreatic α-cells and glucagon granules. To identify αcells, we transduced cells with adenovirus coding for enhanced green fluorescent protein (EGFP) under the control of the pre-proglucagon promoter 66 . The system takes advantage of Tet-On conditional expression in the presence of 4 μM doxycycline, to drive expression of EGFP. The cells were simultaneously transduced with adNPY-mCherry, a well-established secretory granule marker. Alternatively, adenovirus coding for EGFP-tagged neuropeptide Y under control of the preproglucagon promoter (Pppg-NPY-EGFP) was used, thus combining cell type identification and secretory granule label. For both approaches, immunostaining with an anti-glucagon antibody confirmed that over 90% of the fluorescently labeled cells were α-cells ( Fig. 1a and Supplementary Fig 1a). Approximately onethird of the glucagon positive cells were labeled with Pppg-NPY-EGFP (Supplementary Fig 1a). NPY-EGFP labeled granules had excellent overlap with punctate glucagon staining (Fig. 1a), with 94 ± 1% of glucagon positive granules being labeled with NPY-EGFP (37 cells, 5 donors). We verified that exocytosis in the identified cells was stimulated by adrenaline ( Supplementary Fig 1b-d), which increases intracellular Ca 2+ in αbut not β-cells.
Cells were imaged in a standard solution containing (in mM) 138 NaCl, 5.6 KCl, 1.2 MgCl 2 , 2.6 CaCl 2 , 10 D-glucose, 5 HEPES (pH 7.4 with NaOH). In Figs. 3-5 the D-glucose concentration was varied as indicated and cells equilibrated for at least 20 min before recording commenced. Where stated, exocytosis was evoked by elevating K + to rapidly depolarize the cells (75 mM KCl equimolarly replacing NaCl in the standard solution). In these experiments, K + was elevated from 10 s after the onset of the recording until the end of the experiment at 50 s. Timed applications of elevated K + (Figs. 1, 3e, f, 4, 5d black, and 6), adrenalin (Supplementary Figs. 1d and 4), insulin or somatostatin (in Fig. 6), and glucose changes ( Supplementary Fig. 3) were by computer-controlled local pressure ejection from a pulled glass pipette (similar to those used for patch clamp). Spontaneous glucose-dependent exocytosis (Figs. 3a-d, 4a-c, and 6) was recorded for 3 min per after equilibration in the stated conditions. In Fig. 6, insulin or somatostatin were applied during the indicated times.
Image analysis. Exocytosis events were identified based on the characteristic rapid loss of the granule marker fluorescence (1-2 frames). Granule docking events were rare and defined as granules that approached the TIRF field and becoming laterally confined once they reached their maximum brightness 42 .
Docked granules were counted using the 'find maxima' function in ImageJ (http://rsbweb.nih.gov/ij). Values were normalized to each cells' contact area with the coverslip (footprint). The ΔF parameter estimates the fluorescence that is specifically localized to a granule, but subtracting a local background value (average of a 5 pixel wide annulus) from the average fluorescence value in a 3 pixel wide circle, both centered at the granule position.
Immunostaining of pancreatic sections. For analysis of SSTR2 expression, deparaffinized human pancreatic tissue sections (biobank samples obtained from the EXODIAB consortium, Uppsala) were heated in a buffer containing 10 mM Tri-sodium citrate and 0.05% Tween 20 (pH 6) for 15 min, allowed to cool, and rinsed with Dako wash buffer 1x. After a 30-min blocking step (Background Sniper, Biocare Medical), sections were rinsed with wash buffer 1x (Dako) and incubated with anti-SSTR2 (Abcam ab134152, diluted 1:500 in wash buffer), and anti-glucagon antibodies (Dako A0565, diluted 1:1500 in wash buffer) overnight at 4°C. The slides were then washed in wash buffer and incubated with fluorophorelabeled secondary antibodies (diluted in Dako wash buffer 1x) for 30 min at room temperature. Fluorescence was visualized using a Zeiss LSM 780 confocal microscope. For analysis, 3-pixel wide linescans of fluorescent intensity were calculated as illustrated in Fig. 3e, f, top. Background subtracted and estimated as the minimum value to the left of the alignment point (corresponding to the nucleus location), after 3 × 3 median filtering.
Electrophysiology. Standard whole-cell voltage clamp and capacitance recordings were performed using an EPC-9 patch amplifier (HEKA Electronics, Lambrecht/ Pfalz, Germany) and PatchMaster software. Voltage-dependent currents were investigated using an IV-protocol, in which the membrane was depolarized from −70 mV to +80 mV (10 mV steps) during 50 ms each. Currents were compensated for capacitive transients and linear leak using a P/4 protocol. Exocytosis was detected as changes in cell capacitance using the lock-in module of Patchmaster (30 mV peak-to-peak with a frequency of 1 kHz).
During the experiments, the cells were continuously superfused with an extracellular solution containing (in mM) 138 NaCl, 5.6 KCl, 1.2 MgCl 2 , 2.6 CaCl 2 , 10 D-glucose, and 5 Hepes, pH 7.4 adjusted with NaOH at a rate of 0.4 ml/min. All electrophysiological measurements were performed at 32C. In the analysis of the measured voltage-dependent current consists of both Na + and Ca 2+ current components, were the rapid peak current (0-3 ms) represent the Na + current and the sustained current during the latter part of the depolarization reflects the Ca 2+ current. In Fig. 4, for membrane potential, solution was containing (in mM) 76 K 2 SO 4 , 10 KCl, 1 MgCl 2 , and 5 HEPES, pH 7.3 adjusted with KOH.
Statistics. Data are presented as mean ± SEM unless otherwise stated. Statistical significance was tested using t-test for comparing ND and T2D groups, or ANOVA for multiple comparisons, as stated (in Origin 2018). Correlation was quantified as Pearson coefficient r using Excel.
Reporting summary. Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability
The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request. A source data file containing all numeric analysis for Figs. 1c-g, 2b-e, 3b-f, 4b-f, 5a-d, and 6a-h, and Supplementary  Figs. 1d, 2b-d, 3a, b, and 4a is provided with the paper.