ACTAUNIVERSITATIS

UPSALIENSISUPPSALA

2018

Digital Comprehensive Summaries of Uppsala Dissertationsfrom the Faculty of Medicine 1478

α-Cell signalling in glucose-regulated glucagon secretion

QIAN YU

ISSN 1651-6206ISBN 978-91-513-0387-1urn:nbn:se:uu:diva-356478

Dissertation presented at Uppsala University to be publicly examined in B21, BiomedicalCentre, Husargatan 3, Uppsala, Thursday, 13 September 2018 at 09:15 for the degree ofDoctor of Philosophy (Faculty of Medicine). The examination will be conducted in English.Faculty examiner: Professor David W Piston (Department of Cell Biology and Physiology,Washington University School of Medicine, St. Louis, MO, U.S.A).

AbstractYu, Q. 2018. α-Cell signalling in glucose-regulated glucagon secretion. DigitalComprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine 1478.49 pp. Uppsala: Acta Universitatis Upsaliensis. ISBN 978-91-513-0387-1.

Glucagon is a blood glucose-elevating hormone released from α-cells in the islets of Langerhansduring hypoglycaemia. Glucagon is critical for glucose homeostasis and inappropriateregulation of its secretion underlies both impaired counter-regulation of hypoglycaemia andchronic hyperglycaemia in diabetes patients. The mechanisms by which glucose controlsglucagon secretion are poorly understood, but have been suggested to involve both directeffects of the sugar on α-cells and indirect effects mediated by paracrine factors releasedwithin the islet, including insulin and gamma-hydroxybutyrate (GHB) from β-cells, andsomatostatin from δ-cells. This thesis addresses the role of the intracellular messengers ATP,Ca2+ and cAMP in glucose-regulated glucagon secretion. Various fluorescence microscopytechniques were used to monitor changes of these messengers in single, dispersed α-cells andthose in situ within intact islets, and glucagon secretion from islets was measured with animmunoassay. Glucose induced elevations of α-cell ATP, which were smaller and showed a left-shifted concentration-dependence compared to those in β-cells, consistent with α-cells beingless dependent on oxidative metabolism and optimized for sensing hypoglycaemia. α-Cellsshowed Ca2+ oscillations with little glucose dependence. Surprisingly, these oscillations becamesynchronized in phase with Ca2+ oscillations in β-cells at high glucose. Since Ca2+ is a maintrigger of exocytosis in both cell types, and since insulin and glucagon secretion is pulsatile inopposite phase, the results indicate that factors other than Ca2+ are more important for shapingglucagon secretion. Consistent with a key role of cAMP for the regulation of glucagon release,the concentration of the messenger was relatively high in α-cells at low glucose concentrations,and elevations of glucose suppressed cAMP in parallel with glucagon secretion. This effectwas independent of paracrine signalling from insulin and somatostatin. The glucose-inducedsuppression of glucagon secretion was prevented by cAMP-elevating agents and mimickedby inhibitors of protein kinase A. GHB lacked effects both on Ca2+, cAMP and glucagonsecretion from mouse islets, but tended to stimulate glucagon secretion by a somatostatin-receptor-dependent mechanism in human islets. The data indicate that GHB is not an inhibitorof glucagon secretion and that α-cell-intrinsic glucose sensing involves signalling via cAMPand protein kinase A.

Keywords: Islets, glucagon, insulin, somatostatin, ATP, cAMP, Ca , GHB, oscillations, α-cell, β-cell, δ-cell

Qian Yu, Department of Medical Cell Biology, Box 571, Uppsala University, SE-75123Uppsala, Sweden.

© Qian Yu 2018

ISSN 1651-6206ISBN 978-91-513-0387-1urn:nbn:se:uu:diva-356478 (http://urn.kb.se/resolve?urn=urn:nbn:se:uu:diva-356478)

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Dedicated to my dearest family

List of Papers

This thesis is based on the following papers. I. Li, J., Yu, Q., Ahooghalandari, P., Gribble, F.M., Reimann, F.,

Tengholm, A., and Gylfe, E. Sub-membrane ATP and Ca2+ kinetics in -cells: unexpected signaling for glucagon secretion. FASEB J. 2015; 29:3379-3388.

II. Yu, Q., Shuai, H.Y., Ahooghalandari, P., Gylfe, E., and Tengholm, A. Glucose controls glucagon secretion by directly modulating cAMP in α-cells. Submitted manuscript.

III. Yu, Q., Gylfe, E., and Tengholm, A. Quantitative assessment of glucose-regulated cAMP signalling and protein kinase A-mediated glucagon secretion. Manuscript.

IV. Yu, Q., Lai, BK., Ahooghalandari, P., Helander, A., Gylfe, E., Gilon, P., Tengholm, A. γ-Hydroxybutyrate does not mediate glucose inhibition of glucagon secretion. Manuscript.

Reprints were made with permission from the publisher.

Contents

Introduction ................................................................................................... 11 Pancreatic islets ........................................................................................ 12 Glucose-dependence of islet hormone secretion ...................................... 12 Pulsatile release of islet hormones ........................................................... 13 

Stimulus-secretion coupling and oscillatory signalling in β-cells ....... 13 Glucose regulation of glucagon secretion ................................................ 15 

Glucose metabolism in α-cells ............................................................. 15 Neural regulation ................................................................................. 15 Paracrine regulation by β-cell factors .................................................. 15 Paracrine inhibition by somatostatin from δ-cells ............................... 16 Juxtracrine control of glucagon secretion ............................................ 17 α-cell-intrinsic regulation of glucagon secretion ................................. 17 

Role of cAMP in insulin and glucagon secretion ..................................... 19 cAMP production and degradation ...................................................... 19 cAMP effector proteins ........................................................................ 20 Measurements of cAMP ...................................................................... 21 

Aims .............................................................................................................. 22 

Methods ........................................................................................................ 23 Islet isolation, culture, virus infection ...................................................... 23 Measurements of ATP, Ca2+ and cAMP .................................................. 23 Fluorescence microscopy ......................................................................... 25 Measurements of glucagon and insulin secretion ..................................... 26 Signal processing and statistics ................................................................ 27 

Results and discussion .................................................................................. 28 Identification of α- and β-cells within intact islets ................................... 28 Glucose increases [ATP]pm in opposite phase to [Ca2+]pm in α- and β-cells .......................................................................................................... 28 High glucose synchronizes the [ATP]pm and [Ca2+]pm oscillations between α- and β-cells .............................................................................. 29 Somatostatin shapes pulsatile glucagon secretion independently of [Ca2+]pm ..................................................................................................... 30 Dual effects of glucose on cAMP regulation in α-cells ............................ 31 Glucose lowers [cAMP]pm via an α-cell intrinsic pathway independent of paracrine effects of somatostatin and insulin ....................................... 32 

Phosphodiesterase activation contributes to glucose suppression of α-cell [cAMP]pm ....................................................................................... 32 Glucose regulation of α-cell [cAMP]pm does not seem to involve the Ca2+-store operated mechanism ................................................................ 33 cAMP regulation of glucagon secretion is mediated by protein kinase A ............................................................................................................... 33 γ-hydroxybutyrate does not mediate glucose inhibition of glucagon secretion ................................................................................................... 34 

Conclusions ................................................................................................... 36 

Acknowledgements ....................................................................................... 37 

References ..................................................................................................... 39 

Abbreviations

AC Adenylyl cyclase ADP Adenosine diphosphate ATP Adenosine triphosphate [ATP]pm Sub-plasma membrane ATP

concentration [Ca2+]pm Sub-plasma membrane Ca2+

concentration cAMP 3’,5’-cyclic adenosine

monophosphate [cAMP]pm Sub-plasma membrane cAMP

concentration FRET Förster resonance energy transfer GABA γ-aminobutyric acid GHB γ-hydroxybutyrate KATP channel ATP-sensitive K+ channel PDE Phosphodiesterase PFK1 Phosphofructokinase-1 PKA Protein kinase A SOC Store-operated Ca2+ channel SSTR Somatostatin receptor TIRF Total internal reflection fluorescence

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Introduction

Blood glucose homeostasis is maintained mainly by the glucose-elevating and –reducing hormones glucagon and insulin, respectively, released from the islets of Langerhans in the pancreas. When the blood glucose concentration is elevated, such as after a meal, insulin is released into the circulation. Insulin is an anabolic hormone that lowers the production and increases the storage of glucose in the liver by inhibiting glycogenolysis and gluconeogenesis, and by stimulating glycogen synthesis (1). Insulin also promotes glucose uptake and storage in muscle and adipose tissue, and hence causes a lowering of the circulating glucose concentration. Glucagon was first described in 1923 by Kimball and Murlin as a factor that counteracted hypoglycaemia (2), and in 1929, work by Bürger and Kramer demonstrated that this factor increased glucose by a direct glycogenolytic effect on the liver (3). Glucagon is released in response to lowered blood glucose concentrations between meals to enhance the hepatic glucose output by potentiating glycogenolysis and gluconeogenesis and inhibiting glycolysis and glycogenesis (4).

Diabetes is a severe disease characterized by chronic hyperglycaemia. According to the latest IDF report (5), nearly half a billion people suffer from diabetes, and by the end of 2045, the number is expected to reach 629 million. Long-term hyperglycaemia, such as in diabetes, leads to severe and life-threatening health complications, including neuropathy, cardiovascular disease, nephropathy and retinopathy. There are two main forms of diabetes. Type 1 diabetes is usually considered to be caused by an autoimmune attack on the insulin-producing β-cells and requires appropriate daily insulin treatment. The much more common type 2 diabetes, accounting for 90% of all diabetes, is due to insufficient insulin release and resistance to insulin action. The causes of type 2 diabetes are still not well understood but there are strong links to genetics, life style and other environmental factors (5). In many cases, type 2 diabetes can be effectively managed by life-style interventions and pharmacological stimulation of endogenous insulin secretion and action, but severe cases may require exogenous insulin.

While the diabetes treatment almost exclusively is based on restoration of the secretion or action of insulin, the dysregulated glucagon secretion also contributes to the clinical manifestations of the disease. Excessive glucagon levels may aggravate hyperglycaemia (6). Animal experiments have even indicated that increased glucagon levels may be more important than the lack of insulin for hyperglycaemia. Mice devoid of glucagon receptors remain

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normoglycemic despite nearly complete ablation of β-cells, whereas restoration of the expression of glucagon receptors results in severe hyperglycaemia (7,8). Moreover, there is often inadequate glucagon release in response to hypoglycaemia, a frequent and life-threatening complication in insulin-treated patients (6).

Despite the importance of glucagon and the potential benefit of restoring normal glucagon responses in diabetes patients, the mechanisms underlying glucose-regulated glucagon secretion are debated, but consensus is lacking and even fundamental questions remain unresolved. This thesis focuses on intracellular signalling processes in pancreatic α-cells and their importance for glucose regulation of glucagon secretion.

Pancreatic islets The pancreas has both exocrine and endocrine functions. The glucose-regulating hormones are released from clusters of endocrine cells, called islets of Langerhans, which are scattered throughout the exocrine tissue. The islets constitute only 2% of the pancreas volume, but have abundant blood supply, receiving >10% of the blood to the pancreas via the splenic and superior mesenteric arteries. The venous drainage eventually reaches the liver via the portal vein (9). The islets are typically <150 µm in diameter with a cellular composition that varies between species. In rodent islets, approximately 70-80% of the cells are insulin-releasing β-cells, 15-20% are glucagon-releasing α-cells, and up to 10% are δ-cells, which secrete somatostatin, a paracrine factor that regulates secretion from the other islet cells (10,11). There are also a few (<1%) pancreatic polypeptide-releasing PP-cells and ghrelin-secreting ε-cells, which do not seem to be directly involved in blood glucose regulation. The β-cells are clustered in a central core surrounded by a mantle of non-β-cells. Human islets contain fewer β-cells (~50%) and more α-cells (~40%), that are intermingled throughout the islet (10,12). The seemingly random distribution of cells in the human islet should facilitate paracrine interactions between the different cell types (12).

Glucose-dependence of islet hormone secretion Glucose regulates the secretion of all three major islet hormones but with very different concentration dependencies. Insulin secretion is stimulated when the glucose concentration reaches above a threshold of 6-7 mM in mice and ~3 mM in humans, and increases concentration-dependently with half-maximal stimulation at ~11 and maximal stimulation above 20 mM in both species (13). In contrast, glucagon secretion is high at low glucose (0-3 mM) and gradual increase of the glucose concentration inhibits glucagon secretion with

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maximal effect at 5-7 mM for human and mouse islets (14). Further glucose elevation somewhat diminishes this inhibition (14). Somatostatin secretion is stimulated by glucose concentration-dependently throughout the entire range of glucose concentrations up to ~20 mM (14).

Pulsatile release of islet hormones Insulin, glucagon and somatostatin are all released in a pulsatile manner. In healthy individuals this pulsatility results in oscillations of circulating insulin with a periodicity of ~5 minutes (15). The pattern deteriorates in early stages of diabetes and irregular oscillations are found not only in patients with type 2 diabetes (16) but also in first degree relatives (17) and in patients with type 1 diabetes (18). The impaired oscillations may lead to desensitization of insulin receptors, at least in the liver (19), and this process may contribute to hepatic insulin resistance in type 2 diabetes (20,21). Measurements of glucagon secretion from groups of isolated mouse, rat and human islets have shown that glucagon secretion is pulsatile in opposite phase to the insulin pulses at high glucose (22-24). Similarly, measurements of postprandial insulin and glucagon in healthy individuals also revealed an antiparallel relationship between insulin and glucagon oscillations and, notably, this relationship is lost in prediabetic and diabetic patients (25,26). Somatostatin is released in pulses synchronized with those of insulin and, accordingly, in opposite phase to those of glucagon (22,23). The pulsatility of insulin release from islets arises because the secretory activity of β-cells is synchronized via gap junctions (27,28). Such connections do not seem to exist among α-cells or δ-cells, but recent studies have demonstrated the presence of gap junctions between β- and δ-cells (29), which can explain why insulin and somatostatin are secreted in the same phase. It is likely that the inhibitory effects of somatostatin on glucagon secretion contributes to glucagon pulsatility (22,23,30). The synchronization of the many islets in the pancreas required for hormone pulsatility in the portal vein is accomplished by intrapancreatic nerves (31).

Stimulus-secretion coupling and oscillatory signalling in β-cells The pulsatility of islet hormone secretion is thus ultimately driven by β-cells, which individually secrete insulin in a pulsatile manner. At the level of single β-cells, the secretion pattern is determined by cell metabolism and electrical activity. Glucose rapidly enters β-cells via glucose transporters (GLUTs). Rodent β-cells express the high Km GLUT2 transporter whereas human β-cells mainly express GLUT1 with low Km (32). Glucose is subsequently metabolized to generate ATP. The elevated ATP/ADP ratio leads to closure of KATP channels. The resulting membrane depolarization leads to opening of

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voltage-gated Ca2+ channels, influx of Ca2+ and insulin granule exocytosis (33). The cytoplasmic Ca2+ concentration oscillates. There are different types of Ca2+ oscillations. In individual β-cells, a slow pattern (periodicity 2-10 min) is dominating, while in intact islets, there are both slow oscillations and a faster pattern (periodicity 10-60 seconds), as well as combinations of both (34,35). The physiological relevance of the fast pattern, which probably is driven by ionic feedback mechanisms, is unclear, whereas the slow Ca2+ oscillations are believed to generate pulsatile insulin secretion (36). It has been suggested that oscillations in glucose metabolism underlie the slow Ca2+ oscillations (37-39). Indeed, ATP, oxygen consumption, the intracellular concentration of NAD(P)H as well as the mitochondrial membrane potential oscillate with similar slow periodicity as Ca2+ (40-43). The rhythms of these metabolic factors have been attributed to the activity of the glycolytic enzyme phophofructokinase-1 (PFK1). Inhibition of PFK1 activity suppresses oscillatory activity in muscle extracts (39) and it seems likely that PFK1 exerts a similar effect in β-cells. Interestingly, a mutation in this enzyme has been found in human individuals with impaired insulin oscillations (44). The oscillatory activity of PFK1 can be explained by positive feedback from its product fructose 1,6-bisphosphate and even more potent allosteric activation by fructose-2,6-bisphosphate (37), which overcome the inhibitory effects of ATP and citrate (45,46). Metabolic oscillations can occur also independent of glycolysis. For example, citrate oscillations have been observed in isolated β-cell mitochondria (47).

In contrast to the idea that metabolic oscillations drive oscillations of the cytoplasmic Ca2+ concentration, there is evidence that metabolic oscillations do not occur when Ca2+ oscillations are prevented (43,48), suggesting that metabolic oscillations may be generated by Ca2+ oscillations rather than the other way around.

There are several mechanisms by which Ca2+ can influence the cytoplasmic levels of ATP. Elevations of the cytoplasmic Ca2+ concentration stimulates energy requiring transport of Ca2+ into the endoplasmic reticulum (ER) and out of cells via Ca2+ ATPases (49,50). Elevated Ca2+ will also promote uptake of Ca2+ into mitochondria via the mitochondrial Ca2+ uniporter complex (51). This process depolarizes the mitochondrial inner membrane, and hence reduces the driving force for ATP production (42,52,53). However, increased Ca2+ in the mitochondrial matrix stimulates metabolism by activating several dehydrogenases (54,55). Thus, oscillations in metabolism may be shaped by both positive and negative feedback from Ca2+.

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Glucose regulation of glucagon secretion Glucose metabolism in α-cells At variance with mouse β-cells that express the high Km GLUT2, the low Km GLUT1 is the predominant glucose transporter in α-cells (32,56). Although the glucose uptake is relatively slow in GLUT1-expressing cells (57), it is not rate-limiting for the glucose utilization, which instead is determined by glucokinase, the dominant phosphorylating enzyme in both α- and β-cells (32,56,58). Glucose elevation induces a much smaller change of ATP in α-cells than in β-cells (59,60), indicating a lower glucose oxidation and oxidative phosphorylation in α-cells, perhaps due to their high expression of uncoupling protein-2. However, measurements of adenine nucleotides have revealed a 2-fold higher level of ATP in α- than in β-cells at low glucose concentrations (61). Fluorescence imaging of ATP with the ATP sensor Perceval confirmed that elevation of glucose from 1 to 6 mM increases ATP in α-cells from transgenic Glu-RFP mouse (62) but the detailed relationship between glucose, ATP generation and Ca2+ signalling in α-cells still needs to be explored (63).

Neural regulation It has been demonstrated that glucose regulates glucagon secretion via neural control (64). Glucose sensing cells that express GLUT2 have been found in the central nervous system as well as in the hepatoportal vein (65,66). These cells control the activation or inhibition of either sympathetic or parasympathetic neural signals to influence glucagon secretion from the innervated islets (64,67). For example, noradrenaline secreted from the adrenal glands increases glucagon secretion via β2 adrenoceptors on α-cells (64,68). Nevertheless, glucose suppresses glucagon secretion also in isolated islets, indicating that apart from the neural control, α-cell-intrinsic mechanisms and/or paracrine factors are also involved.

Paracrine regulation by β-cell factors Pulsatile glucagon secretion from islets exposed to high glucose concentrations implies that α-cells are synchronized under certain conditions. However, unlike β-cells, which are electrically coupled by gap junctions, α-cells lack such coupling (28) and it is therefore likely that diffusible paracrine factors from neighbouring β- or δ-cells shape pulsatile glucagon secretion. Since glucose stimulates insulin secretion and insulin inhibits glucagon secretion, it has been proposed that insulin from β-cells mediates glucagon suppression by glucose (59,69-71). Accordingly, α-cell-specific insulin receptor knockout in mice results in an enhanced glucagon response to

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hypoglycaemia (72). Not only insulin, but also other factors released from β-cells have been proposed to modulate glucagon secretion. An inhibitory effect of Zn2+ on glucagon secretion in some studies have been suggested to be mediated by activation of α-cell KATP channels and to inhibition of the glycine receptor on α-cells (73,74). Nevertheless, since glucagon secretion is already maximally inhibited below the glucose threshold for stimulation of insulin secretion (75,76), it is unlikely that these β-cell factors regulate glucagon secretion in the hypoglycaemic glucose concentration range.

The inhibitory neurotransmitter GABA, which is generated in β-cells, inhibits glucagon secretion via GABAA-receptor chloride channels (77). Little is known about the regulation of GABA release, but both the content and release of GABA from β-cells is reduced at 20 mM glucose (78), and hence it is difficult to reconcile a role of GABA in the regulation of glucagon secretion in hyperglycaemia. A metabolite of GABA, γ-hydroxybutyrate (GHB), is also an inhibitory neurotransmitter, which has been used for treatment of sleep disorders, narcolepsy and alcohol dependence (79). It has been proposed that glucose stimulates release of GHB from β-cells, and that the transmitter mediates glucose inhibition of glucagon secretion by acting on α-cell GHB receptors (73). The pharmacology of GHB is complex with both low-affinity and high-affinity receptors (79). The most established low-affinity target for GHB is the GABAB receptor, which mediates the effects of millimolar concentrations of exogenous GHB (80). High-affinity receptors have been cloned from mouse and human tissues but their functional profiles remain unclear (81,82). It was recently discovered that the α4β1δ GABAA receptor is a high-affinity GHB target activated by GHB concentrations in the nanomolar to micromolar range (83). The idea that GHB receptor is involved in the regulation of glucagon secretion was based on experiments with ago/antagonists but the direct effect of the neurotransmitter on glucagon secretion has not been investigated.

Paracrine inhibition by somatostatin from δ-cells At variance with insulin, somatostatin is secreted in response to glucose even in the hypoglycaemic concentration range (70,75), making this peptide a better candidate as paracrine mediator of glucose-regulated glucagon secretion. Somatostatin affects both insulin and glucagon secretion via Gi protein-coupled somatostatin receptors (SSTRs). There are five isoforms of SSTRs with SSTR5 and SSTR2 being predominant in mouse β- and α-cells, respectively (84,85). Quantitative PCR data from human islet cells indicate that SSTR2 is predominant in both β- and α-cells (84). Activation of these SSTRs suppresses secretion from α- and β-cells via opening of hyperpolarizing G-protein-gated inwardly rectifying K+ channels, inhibition of cAMP formation and closure of voltage-gated Ca2+ channels (86-88). Nevertheless, although somatostatin potently inhibits glucagon secretion,

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glucagon suppression by glucose persists even in somatostatin knockout mice (89), as well as after perturbing SSTR receptor signalling via SSTR2 blockers or Gαi inhibition (75,90). The available data suggest that paracrine somatostatin signalling exerts tonic inhibition on glucagon secretion but does not mediate inhibition by glucose.

Juxtracrine control of glucagon secretion Yet another model for glucose regulation of glucagon secretion is based on juxtacrine signalling via ephrin and EphA receptors (91,92). Since ephrins, the ligands of EphA receptors, are membrane bound, the ephrin-EphA interaction requires cell-cell contact. By an unknown mechanism, glucose potentiates the forward signalling from β-cell ephrinA ligands to α-cell EphA receptors, which leads to actin polymerization and maintenance of a dense F-actin network in the α-cell. A dense F-actin network prevents exocytosis of glucagon granules and consequently causes suppression of glucagon secretion (91).

α-cell-intrinsic regulation of glucagon secretion Although neural, para- and juxtacrine mechanisms are involved in the regulation of glucagon secretion under certain conditions, it is also evident that glucagon secretion can be regulated via direct α-cell glucose sensing. Such α-cell-intrinsic models have been proposed to explain glucose inhibition of glucagon secretion by closure of voltage-dependent Ca2+ channels with resulting lowering of Ca2+ influx. However, the reduced voltage-dependent Ca2+ influx has been attributed to different mechanisms.

In β-cells, insulin secretion is regulated via modulation of KATP channels by changes of the ATP/ADP ratio. α-Cells also express KATP channels, and they have been attributed a key role in one model for glucose suppression of glucagon secretion (93-96). At low glucose concentrations, most KATP channels in α-cells are closed and this keeps the membrane potential in a range that activates the T-type voltage-dependent Ca2+ channels (VDCCs) and voltage-dependent Na+ channels. The activities of these channels cause additional membrane depolarization and activation of L- and P/Q type VDCCs, which in turn mediate influx of Ca2+ that triggers glucagon secretion (62,93,95,97,98). When the glucose concentration is increased, closure of the remaining KATP channels by the elevated ATP/ADP ratio will slightly depolarize the membrane potential and inactivate the voltage-dependent Na+ channels (97). This inactivation prevents the action potential to reach sufficiently positive values to activate the P/Q-type VDCCs, and the reduced Ca2+ influx leads to inhibition of glucagon secretion (62).

In contrast to this KATP-channel-centred model for glucose-regulated glucagon release, other models are based on α-cell-intrinsic glucose sensing

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that leads to hyperpolarization of the plasma membrane. According to one proposal, glucose regulates glucagon secretion via a Ca2+ store-operated pathway (75,99). Store-operated Ca2+ channels (SOCs) are ubiquitously expressed and their activity is regulated by the interaction with an ER Ca2+-sensor protein, STIM1. Under hypoglycaemic conditions, low energy supply to the α-cell sarco-endoplasmic reticulum Ca2+ ATPase (SERCA) causes depletion of the ER Ca2+ store. Consequently, STIM1 will polymerize and translocate to the plasma membrane to co-cluster with the store-operated channel Orai1, that in turn opens and mediates influx of Ca2+ (100). The opening of SOCs at low glucose depolarizes the membrane to open the L-type VDCCs, influx of Ca2+ and consequently increased glucagon secretion. When the glucose concentration is increased, ATP will stimulate the SERCA pump to refill the ER Ca2+ store, and the STIM1 clusters will disintegrate and the store-operated Ca2+ channels will close. Inactivation of the store-operated channels leads to hyperpolarization, closure of the L-type VDCCs and inhibition of glucagon granule exocytosis secretion (75,99).

Other models propose that glucose hyperpolarizes plasma membrane by energizing the electrogenic Na+ /K+ ATPase or by activating volume-regulated anion channel, that results in inhibition of Ca2+ influx via VDCCs and, hence, suppression in glucagon secretion (101,102). The 2-pore domain K+ channel, TWIK-related acid-sensitive K+ channel, has also been implicated in high glucose-induced hyperpolarization of α-cells and inhibition of glucagon secretion (103).

Fig. 1 Models for intrinsic glucose regulation of glucagon secretion.

See text for details. (A). GLUT1: glucose transporter 1. P/Q type: P/Q type voltage-dependent Ca2+ channel. L-type: L-type voltage-dependent Ca2+ channel. (B). VRAC: volume-regulated anion channel. TASK: TWIK-related acid-sensitive K+ channel. VDCC: voltage-dependent Ca2+ channel. SOC: store-operated Ca2+ channel. Na+ /K+: Na+ /K+ ATPase. SERCA: sarco-endoplasmic reticulum Ca2+ ATPase. ER: endoplasmic reticulum.

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Whereas most models are based on a glucose-stimulated rise of ATP, there is one model that instead assumes that it is the associated drop of AMP at high glucose that inhibits glucagon release via inactivation of the AMP-activated protein kinase (104). This kinase supposedly enhances glucagon secretion independently of Ca2+. There are also other indications that glucose regulates glucagon secretion independently of Ca2+ (105).

Role of cAMP in insulin and glucagon secretion cAMP production and degradation The second messenger cAMP plays a central role in many physiological processes, including insulin and glucagon secretion, where it amplifies Ca2+-dependent exocytosis. The cytoplasmic concentration of cAMP is determined by the rates of production from ATP via adenylyl cyclases (ACs) and degradation by phosphodiesterases (PDEs).

ACs constitute an enzyme family of ten isoforms, nine of which are transmembrane proteins (AC1-AC9) and one which lacks the transmembrane regions and therefore is referred to as soluble AC (106). The transmembrane isoforms share sequence homology with 12 transmembrane domains and two cytosolic regions, which contain the ATP-binding site (107,108). The transmembrane isoforms are primarily activated and inhibited by Gs- and Gi-protein-coupled receptors, respectively. The ACs also show isoform-specific regulation by Ca2+, protein kinases and other factors. For example, AC1 and AC8 are activated, whereas AC3, AC5 and AC6 are inhibited by Ca2+/calmodulin (109,110). Phosphorylation mediated by protein kinase A (PKA) inhibits AC5 and AC6 activity (106), whereas phosphorylation by protein kinase C (PKC) stimulates the Ca2+-insensitive isoforms AC2, AC4 and AC7 but inhibits AC9 (106,111). Many of the ACs are expressed in islets and β-cell lines. For example, the Ca2+-sensitive AC1, AC3 and AC8, as well as the Ca2+-insensitive AC9 have been implicated in the regulation of glucose-stimulated insulin secretion from rodent β-cells. A single nucleotide polymorphism in the gene expressing AC5 shows strong association with type 2 diabetes (112). The soluble AC isoform is regulated by Ca2+ and HCO3

- and probably also by ATP, as it has an ATP affinity in the low millimolar range. Soluble AC has been suggested to account for glucose-induced cAMP production in INS1 cells (113) and mice deficient in this AC isoform has been reported to show reduced glucose-induced cAMP formation and insulin secretion (114). Unlike for β-cells, little information is available about AC expression in α-cells.

The PDEs constitute a large enzyme superfamily terminating cyclic nucleotide signalling by hydrolysing cAMP and cGMP to 5’AMP and 5’GMP. There are 11 structurally related, but functionally distinct, PDE gene families

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(PDE1-11) encoding >50 PDE isoforms (115). PDE3B is the functionally dominating isoform in islets and constitutes >70% of total PDE activity. It is activated by glucose, insulin and cAMP-elevating agent (116,117). Human islets have also been found to express PDE1, PDE4, PDE7, PDE8 and PDE10 (117), several of which have been found to be involved in the regulation of insulin secretion. Much less is known about the expression of PDEs in α-cells, but a recent study suggested that activation of PDE3B is associated with glucose suppression of glucagon secretion (118).

cAMP effector proteins The cellular functions of cAMP are mediated by its effector proteins PKA and Epac. PKA exerts its effect by phosphorylating various proteins. Among substrates relevant for secretion, PKA phosphorylates ion channels, including the KATP channel and VDCCs (119-121). PKA also phosphorylates many proteins involved in the exocytosis machinery such as synaptosomal-associated protein 25 (SNAP-25), synapsin, snapin, and cysteine string proteins, an action that enhances insulin secretion (122). The amplification of secretion by PKA is in part due to sensitization of the exocytosis machinery to Ca2+ (122-125).

Epac is a cAMP-regulated guanine nucleotide exchange factor for Rap proteins, transforming them from an inactive GDP-bound form to the active GTP-bound state. The two isoforms Epac1 and Epac2 are both expressed in pancreatic islets (126,127). Knockout of Epac2 in mice leads to a reduced insulin secretion in response to cAMP elevation, and this effect was suggested to be related to a Rap1-mediated recruitment of granules to the plasma membrane (128). Epac2 also interacts with proteins such as Rim2 (127,129), Piccolo (130) and SUR1 (131) that are important in granule exocytosis.

Glucagon secretion is affected by cAMP-dependent signalling. Adrenaline, which acts via Gs-coupled receptor-mediated cAMP generation, has been found to enhance glucagon secretion (125,132). Large increases of cAMP induced by adrenaline has been proposed to promote glucagon secretion via Epac-dependent activation of L-type Ca2+ channels (125). In contrast, GLP-1, which also generates cAMP, is a well-known inhibitor of glucagon secretion. The inhibitory effect of GLP-1 is likely indirect and mediated by stimulated release of somatostatin (133,134). However, it has also been suggested that GLP-1 has a direct effect on α-cells and that a small rise of cAMP induces PKA-dependent inhibition of P/Q-type Ca2+ channels (98). Glucagon has also been reported to enhance glucagon secretion (135), A recent study using an immunostaining technique has shown that glucose lowers cAMP levels in α-cells and that this effect results from paracrine signalling from insulin and somatostatin. More detailed studies of cAMP regulation in living α-cells are required to clarify if and how the messenger is involved in glucose regulation of glucagon secretion.

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Measurements of cAMP Different fluorescence microscopy approaches have been developed to monitor cAMP dynamics in living cells (136-140). Most biosensors are based on Förster resonance energy transfer (FRET), a non-radiative energy transfer phenomenon first described by Theodor Förster in the 1940s (141). The FRET-based cAMP sensors usually consist of a cAMP-binding protein domain from protein kinase A or Epac, fused to cyan (CFP) and yellow fluorescent protein (YFP) (136,137,139,140). Conformational changes upon cAMP binding results in changed distance between the fluorescent proteins and hence in the degree of FRET. There are also cAMP reporters based on cyclic nucleotide-gated ion channels (142,143). These sensors respond rapidly to cAMP changes, but a potential drawback is that expression of the sensor ion channels may influence cell function via changes in membrane potential and Ca2+ signalling. A third class of cAMP sensors, known as translocation sensors, take advantage of the cAMP-induced dissociation of the regulatory and catalytic subunits of PKA (138). By targeting the regulatory subunit to the plasma membrane, the sensor is particularly well suited for monitoring changes of cAMP in the sub-plasma membrane space, which is the subcellular compartment most relevant for the regulation of exocytosis.

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Aims

The aims of this thesis work were to clarify the mechanisms underlying glucose inhibition of glucagon secretion and the basis for pulsatile glucagon release by:

Clarifying the glucose-dependence of ATP in α-cells and the

relationship between ATP and Ca2+ dynamics in α- and β-cells

Determining how glucose regulates cAMP in α-cells and compare the cAMP levels in α- and β-cells

Elucidate whether alterations in α-cell cAMP underlie glucose-induced changes of glucagon secretion

To clarify if GHB mediates glucose suppression of glucagon

secretion

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Methods

Islet isolation, culture, virus infection Islets of Langerhans were isolated from 5-9 months old C57BL6J mice. The pancreas was excised from the abdominal cavity and then digested with collagenase. After washing, the islets were hand-picked and transferred to RPMI 1640 medium containing 5.5 mM glucose and supplemented with 10% fetal bovine serum, 100 µg/ml penicillin and 100 µg/ml streptomycin, for 1-2 days culture in 37 °C in an atmosphere of 5% CO2. Some experiments were performed after dispersion of the islets to single cells in Cell Dissociation Buffer (Thermo Fisher Scientific) supplemented with 10% (v/v) TrypLE (Thermo Fisher Scientific). The cells were washed by centrifugation and then resuspended in culture medium and seeded on poly-L-lysine-coated 25-mm coverslips at a concentration of ~50,000 cells per glass. After 1-2 days of culture, the islets or cells were infected with adenoviruses encoding the desired fluorescent biosensor constructs and further cultured for 16-18 hours.

Measurements of ATP, Ca2+ and cAMP The changes of ATP, Ca2+ and cAMP concentrations in the sub-plasma membrane space ([ATP]pm, [Ca2+]pm, and [cAMP]pm) were recorded with fluorescent reporters with the signals specifically detected from the membrane and sub-membrane space with total internal reflection fluorescence (TIRF) microscopy (see below).

ATP was recorded with Perceval, a genetically encoded fluorescent reporter consisting of a circularly permuted variant of green fluorescent protein fused to the Mg-ATP-binding bacterial protein GlnK1 (144). Binding of Mg-ATP to GlnK1 induces a pronounced conformation change of the sensor, which leads to increased fluorescence. Perceval was excited at 491 nm and fluorescence emission detected at 527/27 nm (center wavelength/half bandwidth).

To measure changes of the cytoplasmic Ca2+ concentration, islets were in-cubated 30-40 minutes in extracellular buffer containing the acetoxymethyl ester of either of the fluorescent Ca2+ indicators Fura Red or Fluo4. Both indi-cators were excited at 491 nm and their fluorescence emissions were detected at >620 nm and 527/27 nm, respectively.

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For cAMP recordings, islets were infected with adenoviruses encoding either a translocation sensor or a FRET-based reporter. The translocation sensor is composed of a truncated PKA regulatory RIIβ subunit targeted to the plasma membrane with a CAAX motif and a PKA catalytic subunit (Fig. 2A). The membrane-targeted component was either unlabelled or tagged with cyan fluorescent protein (CFP: excited at 445 nm and detected at 483/32 nm) and the catalytic subunit was fused with either yellow fluorescent protein (YFP: excited at 515 nm and detected at 542/27 nm) or the red fluorescent protein mCherry (excited at 561 nm, detected at >620 nm) (138). When the cAMP concentration is low, the PKA catalytic subunit binds to the regulatory subunit at the plasma membrane. When cAMP binds to the regulatory subunit, the catalytic subunit dissociates from the regulatory subunit and is thereby released from the membrane to the cytosol. This translocation can be detected with confocal or TIRF microscopy.

The FRET cAMP biosensor is based on a catalytically inactive and membrane-binding-deficient Epac1 fused to mTurquoise and tandem Venus fluorescent proteins (145). In the absence of cAMP, the two fluorescent proteins are located close enough for FRET to occur between mTurquoise and Venus (Fig. 2B). cAMP binding to Epac1 induces a conformation change that increases the distance between the fluorescent proteins and therefore reduces the degree of FRET. The FRET sensor was excited at 445 nm with emission detected at 483/32 nm (mTurquoise) and 542/27 nm (Venus), respectively. cAMP changes were presented as the ratio of donor fluorescence over the sensitized emission from the acceptor, referred to as the FRET ratio.

Fig. 2 Detection of cAMP with translocation (A) and FRET-based (B) reporters.

See text for details. CFP: cyan fluorescent protein. YFP: yellow fluorescent protein. RIIβ: Truncated PKA regulatory subunit RII. Cα: PKA catalytic subunit. mTur:

mTurquoise. VEN: Venus. REM: Ras exchange motif in the catalytic region of Epac1. GEF: a catalytically inactive guanine nucleotide exchange factor domain of Epac1. VLVLE: a conserved motif of the cAMP-binding domain. CNB: cyclic nu-cleotide binding domain.

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In study III, attempts were made to calibrate the FRET ratio to record ab-solute intracellular cAMP concentrations. This was made by exposing perme-abilized cells expressing the FRET reporter to known concentrations of cAMP. The cells were superfused with intracellular-like medium containing (in mM) 140 KCl, 6 NaCl, 0.4 CaCl2, 1 MgCl2, 10 HEPES, 2 EGTA, 1 MgATP, 0.1 IBMX with pH adjusted to 7.00 with KOH. The superfusion was stopped and α-toxin added to the chamber to permeabilize the plasma mem-brane. α-toxin allows passage of ions and small molecules, whereas proteins are retained inside the cells. After 15 minutes treatment with the toxin, super-fusion was restarted. cAMP was subsequently added in increasing concentra-tions (1-3000 µM) to establish the relationship between cAMP concentration and the FRET response.

cAMP concentration changes were recorded as the FRET ratio. The FRET ratios were plotted against the cAMP concentration and the data fitted to a sigmoidal curve described by the Hill equation using Igor Pro software (Wavemetrics, Lake Oswego, OR, USA). The curve was used to determine the parameters rate (describing the steepness of the curve) and half-max (the cAMP concentration yielding half-maximal FRET response) required for the translation of FRET responses to absolute cAMP concentration. The maximum and minimum FRET ratios are also required and these parameters were estimated in intact cells during exposure to the combination of 10 µM forskolin and 100 µM IBMX (maximal FRET ratio at saturating cAMP, Rmax) and during exposure to the adenylyl cyclase inhibitor SQ22536 (minimum detection level, Rmin). The cAMP concentration was then calculated according to the following equation:

10

Fluorescence microscopy Before imaging, the islets or cells were preincubated for 30-40 min in an extracellular experimental buffer containing (in mM) 4.8 KCl, 125 NaCl, 1.28 CaCl2, 1.2 MgCl2, 3 or 7 glucose, 25 HEPES with pH adjusted to 7.40 with NaOH. Islets were then allowed to attach on a poly-L-lysine coated coverslip. The coverslips with islets or cells were used as bottoms of a 50-µl (objective TIRF) or 140-µl (prism TIRF) microscopy chamber and superfused with experimental buffer at a rate of 200 or 350 µl/min thermostated at 37 °C.

Most imaging experiments were performed with TIRF microscopy. Two different microscope setups were used. An objective-based TIRF system was used for experiments with intact islets. Laser light was coupled to a TIRF

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illuminator attached to an inverted Nikon Eclipse Ti microscope with a 60x, 1.45-NA oil immersion objective. A prism-based TIRF setup was used for single-cell experiments. It is based on a Nikon E600FN upright microscope with a 10x, 0.3-NA water immersion objective. In the prism-based setup, the laser beam was homogenized and expanded by a rotating light shaping diffuser and refocused through a modified quartz dove prism (Axicon, Minsk, Belarus) with a 70º angle to achieve total internal reflection. Some experiments with FRET-based cAMP sensor was performed with a confocal microscope based on a spinning-disk confocal scan head (Yokogawa CSU-10) attached to a Nikon microscope with a 100x, 1.49-NA oil immersion objective. All setups were equipped with diode or diode-pumped solid-state lasers for excitation at 445, 491, 515 and 561 nm (Cobolt, Stockholm, Sweden). Emission light was selected with interference filters mounted in a filter wheel (Lambda 10-3, Sutter instruments). Images were acquired with a back-illuminated EMCCD (electron-multiplying charge-coupled device) camera (DU-888, Andor Technology, Belfast) controlled by Metafluor software (Molecular Devices, Downingtown, PA). Image analysis were with Metafluor software.

Measurements of glucagon and insulin secretion In study I, batches of size-matched islets (8-10) were preincubated in experimental medium containing 3 mM glucose for 30 minutes and then transferred to new medium containing different glucose concentrations and test substances and incubated for another 40 minutes. The incubation medium was collected for measurements of secreted hormones while the islets were sonicated in acid ethanol for determination of total hormone content. Insulin was measured with an immunoassay kit from Mesoscale Discovery (Rockville, MD) and glucagon was measured with an ELISA kit from Mercodia AB (Uppsala, Sweden).

Most secretion experiments were performed using a slow perfusion setup. Groups of 12-15 islets were loaded into a sealed Teflon tube. The tube was thermostated at 37 °C in a water bath and perfused with experimental medium at a rate of 0.06 ml/min. After 30 minutes’ equilibration in 7 mM glucose, the perfusate was collected every 5 minutes and 3-5 fractions were collected at each experimental condition. Glucagon was measured with ELISA and secretion was expressed as the average of the last two fractions at each condition. A similar setup was used to determine the kinetics of hormone secretion, but the islets were instead perifused at 150 µl/min. The perifusates were collected every 40 s or 90 s and analysed with ELISA.

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Signal processing and statistics Statistic comparisons between groups were assessed with student’s t-test or one-way ANOVA followed by post hoc paired t test with the Holm-Bonferroni sequential correction. Asymmetric sliding window cross correlation was performed in Study I to assess the correlation between two signals. This analysis was performed with MATLAB (The Mathworks, Natick, MA, USA).

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Results and discussion

Identification of α- and β-cells within intact islets The understanding of α-cell signalling has been hampered by difficulties to identify these cells among more abundant β-cell within intact islets. In order to study α-cells within the natural islet environment, feasible and reliable criteria to identify the different cell types in islets are required. For this purpose, we used Glu-RFP transgenic mice expressing red fluorescent protein specifically in α-cells to establish functional criteria for their identification. In the dominating RFP-negative β-cells, [Ca2+]pm was low and stable at 3 mM glucose and elevation of the concentration to 20 mM induced an initial lowering of [Ca2+]pm followed by pronounced elevation with oscillations, consistent with previous reports (146,147). In contrast, the RFP-positive α-cells that usually had smaller footprints under TIRF microscopy, showed irregular [Ca2+]pm spiking in the low glucose concentration range while 20 mM often but not always temporarily inhibited the spiking, which resumed after ~10 min. Introduction of glutamate induced marked increase of [Ca2+]pm in the α-cells but had little effect on the β-cells, consistent with expression of functional ionotropic glutamate receptors only in the α-cells (148). Thus, based on the size of the cellular footprints and [Ca2+]pm signalling at low and high glucose as well as in response to glutamate, we could discriminate between α- and β-cells within intact islets. Moreover, it is unlikely that somatostatin-releasing δ-cells or pancreatic polypeptide-secreting PP-cells were mistaken as α- or β-cells, since they are rarely observed when studying the superficial islet cell layer with TIRF microscopy (149).

Glucose increases [ATP]pm in opposite phase to [Ca2+]pm

in α- and β-cells ATP has been attributed important messenger functions mediating both glucose-regulated insulin and glucagon secretion. However, whereas glucose-generated ATP stimulates insulin secretion the release of glucagon is inhibited. Since the underlying ATP dynamics in α-cells is unknown we used the ATP biosensor Perceval to compare the effects of glucose on [ATP]pm in islet α-cell and β-cells. Elevating glucose from 1 to 5 mM induced comparable [ATP]pm increases in α- and β-cells whereas further elevation to 20 mM

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elicited a much smaller [ATP]pm response in the α- than in the β-cells. These data support the previous view that α-cells have lower glucose oxidation rate (61,150) and perhaps less efficient oxidative phosphorylation due to the high expression of uncoupling protein 2 (151). However, when expressed in relation to the effect of 20 mM glucose the relative [ATP]pm increase induced by 5 mM of the sugar was higher in α-cell than that in β-cells, indicating a left-shifted dose-response relationship consistent with the expectation that α- and β-cells are primarily responsive to hypo- and hyperglycaemia, respectively. This finding adds to previous static measurement of adenine nucleotides indicating that α-cell ATP reaches a maximum already at low glucose concentrations (61).

Since ATP is required for Ca2+ transport and Ca2+ may have both positive and negative effects on mitochondrial ATP production (42,49,50,52-55), it is important to clarify the crosstalk between the two messengers. The ATP/ADP ratio directly affects the plasma membrane KATP channels, which have been implicated in the regulation of both insulin and glucagon secretion. The relationship between Ca2+ and ATP in the sub-plasma membrane space of α- and β-cells within pancreatic islets was therefore studied. Consistent with previous data from our group (152), 20 mM glucose induced regular oscillations of [ATP]pm and [Ca2+]pm that were in opposite phase in the β-cells. The individual α-cells within the same islets showed irregular oscillations of both messengers at 3 mM glucose, which were often temporarily inhibited by 20 mM and then resumed. Also the α-cell [ATP]pm and [Ca2+]pm oscillations were in opposite phase. These data indicate that in both cell types, ATP consumption for elimination of Ca2+ from the cytoplasm, and potentially inhibitory effects of Ca2+ on mitochondrial ATP production, are more important determinants of [ATP]pm than any positive effect of Ca2+ on ATP production.

High glucose synchronizes the [ATP]pm and [Ca2+]pm

oscillations between α- and β-cells The β-cells showed low and stable [Ca2+]pm and [ATP]pm at 3 mM glucose, and the oscillations of [Ca2+]pm and [ATP]pm in opposite phase induced by 20 mM of the sugar became almost perfectly synchronized among the different islet β-cells due to their coupling by gap junctions (27,28), which is a prerequisite for the pulsatile release of insulin. However, the irregular [Ca2+]pm oscillations with [ATP]pm oscillations in opposite phase that were observed in individual α-cells exposed to 3 mM glucose lacked apparent synchronization with other islet α-cells, which is consistent with lack of gap junction coupling of these cells (28) and steady non-pulsatile release of glucagon under low glucose conditions (22,23). The circulating insulin and glucagon

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concentrations oscillate in opposite phase in normal subjects (26), a phenomenon apparently reflecting pulsatile insulin and glucagon release in opposite phase from the pancreatic islets that has been documented both with glucose-stimulated human (23) and mouse (22) islets. It was therefore expected that [Ca2+]pm, which is believed to initiate both insulin and glucagon secretion, should oscillate in opposite phase in α- and β-cells. Nevertheless, 20 mM glucose, which induces pulsatile release of insulin and glucagon in opposite phase, tended to synchronize the [Ca2+]pm oscillations among the gap junction-coupled β- and non-coupled α-cells into the same phase. The synchronization between β- and α-cells and among the α-cells was not as perfect as that between the coupled β-cells but became striking when averaging [Ca2+]pm among the α- and β-cells, respectively, within each islet. Likewise, the [ATP]pm oscillations, which were in opposite phase to [Ca2+]pm

oscillations in both α- and β-cells, also tended to synchronize between α- and β-cells at high glucose. Such gap junction-independent synchronization may depend on diffusible paracrine factors secreted from β- or δ-cells within the same islets in high glucose. ATP for example, which is co-released with insulin, has been reported to coordinate the rhythmicity of the glucose-induced [Ca2+]pm oscillations among dispersed β-cells that lack physical contact (153,154). However, the enigma remains, why is pulsatile release of insulin and glucagon in opposite phase when the secretion-triggering Ca2+signal tends to synchronize in the same phase between α- and β-cells?

Somatostatin shapes pulsatile glucagon secretion independently of [Ca2+]pm

Since pulsatile glucagon secretion is generated in parallel with pulsatile insulin release we hypothesized that the opposite phase between the hormone pulses is generated by some paracrine factor that inhibits glucagon secretion even when α-cell [Ca2+]pm is high. Somatostatin is released in a pulsatile manner from the δ-cells in the same phase as the insulin pulses (22,23) and both hormones have been suggested to mediate glucose inhibition of glucagon secretion (59,69-71,155). We therefore explored the possible involvement of these hormones in the generation of pulsatile glucagon release. The insulin receptor antagonist S961 neither affected [Ca2+]pm in α- or β-cells, nor insulin or glucagon secretion. On the contrary, a somatostatin receptor 2 antagonist potently increased glucagon secretion at both 3 and 20 mM glucose but had little effect on the [Ca2+]pm oscillations in α-cells. Somatostatin but not insulin is therefore a strong candidate acting downstream of [Ca2+]pm to generate pulsatile glucagon secretion in opposite phase to insulin and somatostatin when secretion of the latter hormones is stimulated. The synchrony between pulsatile insulin and somatostatin secretion is not unexpected since the

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[Ca2+]pm oscillations are synchronized between islet β- and δ-cells (149), which may involve both paracrine β-cell factors like ATP (153,154) and direct gap junction coupling between these cell types (29).

Although somatostatin has pronounced effects on glucagon secretion, glucose inhibition of glucagon secretion was neither prevented in the presence of SSTR2 receptor antagonist nor after blocking any Gi-mediated inhibitory effect of somatostatin with pertussis toxin pre-treatment (90). Moreover, glucose concentrations that strongly inhibit glucagon secretion often failed to affect [Ca2+]pm in α-cells or only induced a temporary suppression. These data indicate that additional mechanisms are required to explain glucose regulation of glucagon secretion, especially under hypoglycaemia conditions when the hormone mediates glucose counterregulation under conditions with low release of paracrine factors from non-α-cells.

Dual effects of glucose on cAMP regulation in α-cells cAMP is an important second messenger amplifying the triggering effect of Ca2+ on insulin secretion (156). Since Ca2+ also triggers glucagon granule exocytosis (125), and cAMP-elevating stimuli potently augment hormone release (132,135) it has been assumed that the messengers have similar roles in the two cell types. ATP is the immediate precursor of cAMP and since we found that [ATP]pm was relatively more elevated by low glucose concentrations in α- than in β-cells, we hypothesized that cAMP might also be elevated in α-cells under such conditions. Indeed, in islets exposed to only 1 mM glucose, an adenylyl cyclase inhibitor induced pronounced lowering of [cAMP]pm in the α-cells, whereas the concentration in β-cells was too low to detect any further reduction. To more precisely pinpoint the concentration differences, we took advantage of a FRET-based cAMP reporter that was calibrated with different cAMP concentrations in α-toxin permeabilized dispersed islet cells. The permeabilization approach could not be applied to cells within islets but the calibration curves obtained with free cells showed little variation in slope and half-maximal response. These data were therefore used together with in situ estimates of minimal and maximal cAMP responses to estimate the absolute concentration. This approach confirmed that at 1 mM glucose, [cAMP]pm was much higher in α- than in β-cells. Supporting the importance of glucose-generated α-cell ATP for the cAMP formation, we found that [cAMP]pm increased with glucose concentration in the 0-1 mM concentration range. More importantly, inhibition of glucagon secretion by further elevation of glucose to 7 mM was associated with pronounced reduction of [cAMP]pm in the α-cells. Based on these data, we argued that in the hypoglycaemic concentration range, glucose may suppress glucagon secretion by lowering cAMP. Lowering of cAMP has previously been suggested to mediate glucose-induced paracrine inhibition of glucagon

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secretion by somatostatin and insulin (118). Considering that the paracrine influence is expected to be minimal in hypoglycaemia it is important to investigate whether glucose can also control cAMP via α-cell-intrinsic mechanisms.

Glucose lowers [cAMP]pm via an α-cell intrinsic pathway independent of paracrine effects of somatostatin and insulin To clarify whether the lowering of α-cell [cAMP]pm by glucose elevation involves other mechanisms than paracrine release of insulin or somatostatin from the neighbouring β- and δ-cells, the signalling of these hormones was inhibited with receptor antagonists. Addition of insulin or the insulin receptor antagonist S961 did not affect α-cell [cAMP]pm at either low or high glucose and S961 had no effect on glucagon secretion. In contrast, somatostatin suppressed α-cell [cAMP]pm markedly at 1 mM glucose. However, although somatostatin receptor 2 antagonism increased α-cell [cAMP]pm and stimulated glucagon release, it neither prevented the glucose-induced [cAMP]pm lowering in the α-cells nor the accompanying inhibition of glucagon from pancreatic islets. Further evidence for a somatostatin-independent effect was obtained with the observation that the glucose dependent [cAMP]pm changes in the α-cells persisted after blocking any Gi-mediated inhibitory effect of somatostatin with pertussis toxin pre-treatment or minimizing the paracrine effects by omitting Ca2+. These data support previous findings that the paracrine role of somatostatin in glucagon secretion is more to mediate tonic inhibition than the inhibitory effect of glucose (75,89,90). Our data consequently indicate that the α-cells have intrinsic mechanisms by which variations in the glucose concentration translate into changes of cAMP.

Phosphodiesterase activation contributes to glucose suppression of α-cell [cAMP]pm We next investigated whether the α-cell-intrinsic lowering of [cAMP]pm by glucose elevation could be attributed to inhibition of cAMP-generating ACs or activation of cAMP degradation by PDEs. Dispersed α-cells or islets were therefore exposed to an intermediate concentration of the general PDE inhibitor IBMX to elevate cAMP without saturating the sensor. Some dispersed α-cells subsequently responded to a 7 to 1 mM glucose lowering with a smaller cAMP increase than in the absence of IBMX as expected if glucose were activating PDEs. In one cell, the cAMP response to glucose reduction was instead enhanced as if the cAMP change were due to activation

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of ACs. However, in most α-cells there was no effect of IBMX on the [cAMP]pm response to glucose reduction. Among α-cells within intact islets, a higher proportion (28%) showed an enhanced [cAMP]pm elevation in response to glucose reduction in the presence of IBMX, maybe because of the paracrine influence from somatostatin. Nevertheless, in the majority of the islet α-cells, IBMX reduced the cAMP response to glucose lowering consistent with glucose-mediated regulation of PDE activity.

Glucose regulation of α-cell [cAMP]pm does not seem to involve the Ca2+-store operated mechanism Previous studies have implicated a role of the Ca2+ store-operated Ca2+ influx mechanism in α-cell-intrinsic glucose regulation of glucagon secretion (75,99). Since the store-operated mechanism has also been found to control cAMP formation (157) we investigated its possible involvement in glucose regulation of α-cell cAMP. The results were not entirely clear. In support for a role of the store-operated mechanism, inhibition of Ca2+ influx through the SOCs with 2-APB markedly reduced the cAMP elevation induced by glucose reduction. On the other hand, SOC activation by emptying ER with the SERCA pump inhibitor CPA enhanced the glucose-induced cAMP changes, indicating that glucose reduction-induced cAMP elevation is not mediated by SOC activation. Since 2-APB is also an inhibitor of IP3 receptors and modulator of transient receptor potential channels, its effect on cAMP regulation may be mediated by other mechanisms than store-operated Ca2+ entry.

cAMP regulation of glucagon secretion is mediated by protein kinase A Since glucagon has been found to elevate α-cell cAMP (135) it is important to clarify whether the glucose-induced changes in α-cell cAMP determine glucagon secretion or if they simply result from an autocrine effect of the released hormone. Our data strongly support that cAMP controls glucagon secretion since inhibition of hormone release by glucose elevation was abolished when [cAMP]pm was clamped at a high level by exposure to the phosphodiesterase inhibitor IBMX or a membrane-permeable cAMP analogue.

Our quantification of α-cell cAMP indicated that the concentration was above the threshold for activation of PKA (158) when glucagon release was stimulated by 1 mM glucose. Further support that PKA mediates the effect of cAMP was obtained by addition of the PKA inhibitors Rp-8-CPT-cAMPS or

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Myr-PKI at 1 mM glucose, which mimicked the inhibitory effect of glucose elevation on glucagon secretion. Taken together, these results indicate that stimulation of glucagon secretion in response to hypoglycaemia depends on cAMP elevation and signalling via PKA.

γ-hydroxybutyrate does not mediate glucose inhibition of glucagon secretion Based on studies with GHB receptor ago-/antagonists and the observation that glucose stimulates GHB release from islet β-cells, this GABA metabolite has been proposed to mediate glucose-induced paracrine suppression of glucagon release (73). However, direct observations with GHB, which is a drug-classified compound with restricted availability, are still lacking. We now tested GHB and found that it had little effects on Ca2+ or cAMP signalling in mouse α-cells, and that it didn’t affect glucagon secretion at either 1 or 7 mM glucose. Moreover, the GHB receptor ago- and antagonists used in the previous study had little effect on insulin and glucagon secretion from mouse islets. However, GHB surprisingly stimulated glucagon secretion from human islets. Since GHB is believed to be an inhibitory neurotransmitter (79) we tested whether its stimulatory effect might be mediated by suppression of somatostatin secretion. Indeed, GHB stimulation of glucagon release was prevented in the presence of a SSTR2 receptor antagonist. These data strongly indicate that GHB is not a paracrine mediator of glucose-inhibited glucagon secretion from α-cells but indicate that human δ-cells have receptors binding GHB, the physiological significance of which remains to be determined.

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Fig 3. Model of glucose regulation of glucagon secretion from α-cells.

Glucagon secretion depends on Ca2+ influx though VDCCs. Different models have been proposed to explain glucose regulation of glucagon release by modulation of the channel activity. The present thesis instead suggests that Ca2+ has a permissive role and that the major effect of glucose is to control cAMP, which is a strong amplifier of glucagon exocytosis. In hypoglycaemia, the α-cell senses the glucose reduction and generate more cAMP to stimulate glucagon release and restore normal glycaemia. This effect seems to involve inhibition of cAMP-degrading phosphodiesterases and perhaps also activation of cAMP-generating adenylyl cyclases. In hyperglycaemia, paracrine mechanisms generate pulsatile release of glucagon in opposite phase to the pulsatile release of insulin and somatostatin. This scenario is ultimately driven by the dominating gap junction-coupled β-cells in which high glucose induces Ca2+ oscillations in phase with pulsatile insulin release. The β-cells also synchronize Ca2+ signalling and pulsatile secretion of somatostatin from the δ-cells by direct coupling via gap junctions. Also Ca2+ signalling in the α-cells tend to synchronize with that in the β-cells, probably via release of ATP. However, the inhibitory effect of somatostatin is stronger than the stimulatory action of Ca2+ explaining the generation of pulsatile glucagon secretion in opposite phase to the average changes in α-cell Ca2+.

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Conclusions

1. α-Cells have smaller [ATP]pm responses to increasing glucose concentrations than β-cells but are relatively more sensitive to glucose in hypoglycaemia.

2. Glucose induces [ATP]pm and [Ca2+]pm oscillations in opposite phase in both α- and β-cells, reflecting ATP consumption for Ca2+ removal from the cytoplasm.

3. Glucose concentrations that stimulate insulin release tend to synchronize oscillatory [Ca2+]pm signalling between α- and β-cells although pulsatile release of glucagon and insulin is in opposite phase. The [Ca2+]pm synchronization of the α-cells is probably mediated by paracrine influences from the electrically coupled β-cells.

4. Pulsatile release of glucagon in opposite phase with that of insulin is generated by pulsatile release of somatostatin from the δ-cells paralleling the pulsatile β-cell secretion of insulin. The inhibitory effect of somatostatin on glucagon secretion apparently dominates over the stimulation expected from α-cell Ca2+ elevation.

5. [cAMP]pm in the α-cells increases with glucose concentration in the

0-1 mM, probably reflecting generation of the cAMP precursor ATP. Further glucose elevation instead lowers [cAMP]pm by an α-cell-intrinsic mechanism that probably involves PDE activation. At hyperglycaemic glucose concentrations, paracrine factors like somatostatin likely contribute to lower α-cell cAMP.

6. Glucose regulation of glucagon secretion is mediated by changes in cAMP and PKA activity rather than by Ca2+, which seems to have a more permissive role.

7. GHB does not mediate glucose-suppressed glucagon secretion, but stimulates glucagon secretion from human islets by paracrine inhibition of somatostatin release.

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Acknowledgements

This thesis was carried out at the department of Medical Cell Biology, Uppsala University, Sweden. Here I would like to express my sincere gratitude to all the people that helped and supported me during the thesis work: Great thanks to my main supervisor Anders Tengholm for your guidance, support and fruitful advices both in work as well as my life. You are always the first person I turned to when I faced different decisions in work and life. Thank you for believing me and choosing me as a PhD student in your group. Thank you for helping me to balance my PhD work time in Sweden and my family time in U.S. I was always encouraged by your passion and enthusiasm in science and indeed learned a lot in different aspects from you. I really had a great time in our lab and you are the most important reason why I loved Uppsala. Sincere thanks to my co-supervisor Erik Gylfe for your inspiration, patience and accepting me to do the master thesis in this nicest lab. I am so lucky to have a supervisor that cares so much about my project, responds to my ques-tion and continuously supports me in my studies. Your wise ideas, suggestions as well as the warmest smile really guide me throughout the whole thesis work. Thanks to my co-supervisor Sebastian, for always sharing your vast knowledge and nice suggestions during my thesis work. Your critical com-ments made the thesis better. Thanks to all my lovely current and formal lovely group members. Olof for always being the go to person when I got scientific questions during my thesis work as well as all the funny chats in the lab. Beichen, for all the nice time spending with you and supplying numerous nice food when I was hungry. Jia, for always being there anytime when I got problems and all the fun time with you. Hongyan, Yunjian, Mingyu and Ida, for all happy office time and help in my life and studies. Antje, My and Nadine, for the interesting knitting and baking workshop. Special thanks to Parvin, for performing important exper-iments for this thesis as well as for always caring me. Bo, Eva, Helene, Oleg and Gonzalo, for wonderful suggestions and making a so enjoyable lab envi-ronment.

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Thanks to all the colleagues in the department for helping me with different problems and creating a nice work atmosphere. The head of the department: Professor Nils Welsh. Thanks to my dear friends in Sweden for sharing happy time with me. Thanks to the Friday fika group for the wonderful cakes and chatting. Finally, great thanks for my dear parents for their endless love. My dear hus-band Lei Cheng for your great understanding and things we have been through. I love you!

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A doctoral dissertation from the Faculty of Medicine, UppsalaUniversity, is usually a summary of a number of papers. A fewcopies of the complete dissertation are kept at major Swedishresearch libraries, while the summary alone is distributedinternationally through the series Digital ComprehensiveSummaries of Uppsala Dissertations from the Faculty ofMedicine. (Prior to January, 2005, the series was publishedunder the title “Comprehensive Summaries of UppsalaDissertations from the Faculty of Medicine”.)

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