Role of the type II diabetes- associated gene SLC30A8 in ...€¦ · 4 Abstract The SLC30A8 locus...
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Role of the type II diabetes- associated gene SLC30A8 in the pancreatic α-cell Antonia Solomou Imperial College London Department of Medicine Division of Diabetes, Endocrinology & Metabolism Section of Cell Biology
Transcript of Role of the type II diabetes- associated gene SLC30A8 in ...€¦ · 4 Abstract The SLC30A8 locus...
Role of the type II diabetes-
associated gene SLC30A8 in
the pancreatic α-cell
Antonia Solomou
Imperial College London
Department of Medicine
Division of Diabetes, Endocrinology & Metabolism
Section of Cell Biology
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DECLARATION OF ORIGINALITY I declare that the following work presented in this thesis is original, except where
indicated by special reference in the text, and that no part of the dissertation has
been submitted for any other degree.
Some of the results contained here, have been presented at conference and
seminars and have been submitted to the scientific literature for publication.
COPYRIGHT DECLARATION
The copyright of this thesis rests with the author and is made available under a
Creative Commons Attribution Non-Commercial No Derivatives licence. Researchers
are free to copy, distribute or transmit the thesis on the condition that they attribute
it, that they do not use it for commercial purposes and that they do not alter,
transform or build upon it. For any reuse or redistribution, researchers must make
clear to others the licence terms of this work.
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Acknowledgements
First, I would like to thank Prof Rutter for giving me this opportunity to work in the
lab and earn a PhD. His knowledge and guidance were much appreciated. Secondly, I
would like to thank Dr Hodson for his patience and invaluable help throughout my
PhD, especially in the microscope room.
Life in the lab would have been a lot more boring without Matthew, Ryan, Sophie,
Nick and Natalie. Thanks for all the laughter and the excessive amounts of biscuits
we shared. To all the other lovely people in the group, many of whom have helped
me throughout the years; it has been great working in such a pleasant and relaxed
environment.
Last but certainly not least, I would like to thank Tom, my love and future husband,
who has been there for me right from the start with his love and support whenever I
needed it.
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Abstract
The SLC30A8 locus encodes the Zn2+ transporter ZnT8, whose expression is largely
restricted to α- and β-cells, endocrine cells of the pancreatic islet. Genome-wide
association studies have revealed >90 loci associated with Type 2 diabetes. The first
such study identified a specific single nucleotide polymorphism that results in an
amino acid substitution and a transporter with reduced activity. This risk variant is
associated with increased Type 2 diabetes risk. More recently, rare loss-of-function
variants were found to be protective. Here, we aimed to investigate the role of
SLC30A8/ZnT8 in the regulation of glucagon secretion. ZnT8 was selectively deleted
in the α-cell by crossing mice bearing floxed alleles at exon 1 with mice carrying a Cre
recombinase transgene under the control of the preproglucagon promoter.
Additionally, these mice were crossed to Rosa26 RFP mice for identification of α-
cells. Fluorescence-activated sorting of RFP+ cells revealed that recombination at the
RFP locus occurred in ~30% of α-cells. ZnT8 was deleted in ~50% of these labelled
cells achieving a total of ~15% deletion in the whole α-cell population. Glucose
tolerance and insulin sensitivity were normal, though during hypoglycaemic clamps
female KO mice required lower glucose infusion rates and had enhanced glucagon
secretion compared to control mice. Similarly, isolated islets from KO mice released
significantly more glucagon at 1mM glucose. In addition, inducible overexpression of
ZnT8 led to a reciprocal decrease of glucagon secretion from isolated islets at 1mM
glucose. Hypoglycaemic clamps confirmed the finding of lowered glucagon release
following ZnT8 overexpression. Cytosolic Ca2+ responses to low glucose were
comparable in WT and KO cells. Both cytoplasmic and granular free Zn2+ levels were
significantly reduced in the KO α-cells compared to control cells yet there were no
changes in the gene expression levels of other ZnT famlily members. Thus it appears
that ZnT8 is important in the α-cell for appropriate responses to hypoglycaemia and
Zn2+ homeostasis.
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Table of contents
DECLARATION OF ORIGINALITY / COPYRIGHT DECLARATION .................................................. 2
Acknowledgements ................................................................................................................... 3
Abstract ..................................................................................................................................... 4
Table of contents ....................................................................................................................... 5
List of figures ........................................................................................................................... 10
Abbreviations .......................................................................................................................... 11
Chapter 1 - Introduction ......................................................................................................... 14
1.1 The pancreas ................................................................................................................. 15
1.2 Islets of Langerhans ....................................................................................................... 15
1.2.1 Islet architecture .................................................................................................... 15
1.2.2 Islet microvasculature ............................................................................................ 16
1.2.3 Islet innervation ...................................................................................................... 17
1.3 Glucagon ........................................................................................................................ 19
1.3.1 Historical perspectives ........................................................................................... 19
1.3.2 Pancreatic α-cell development ............................................................................... 19
1.3.3 Preproglucagon – transcriptional control and processing ..................................... 20
1.3.4 Glucagon receptor .................................................................................................. 24
1.3.5 Physiological actions of glucagon ........................................................................... 24
1.4 Glucagon secretion ........................................................................................................ 25
1.4.2 Regulation by glucose: paracrine or intrinsic? ....................................................... 27
1.4.3 Kinetics of exocytosis.............................................................................................. 28
1.4.4 Stimulation of exocytosis ....................................................................................... 29
1.5 Metabolism of the α-cell ............................................................................................... 30
1.5.1 Glucose ................................................................................................................... 30
1.5.2 Amino acids ............................................................................................................ 32
1.5.3 Fatty acids ............................................................................................................... 33
1.6 Regulators of glucagon secretion – autocrine, paracrine and endocrine ..................... 33
1.6.1 Insulin ..................................................................................................................... 34
1.6.2 Zinc ......................................................................................................................... 35
1.6.3 Somatostatin .......................................................................................................... 35
1.6.4 GABA ....................................................................................................................... 36
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1.6.5 Glutamate ............................................................................................................... 36
1.6.6 Glucagon-like-peptide (GLP-1) ............................................................................... 37
1.6.7 ATP .......................................................................................................................... 38
1.6.8 Glucagon ................................................................................................................. 38
1.6.9 Ghrelin .................................................................................................................... 39
1.7 Neural regulation of glucagon secretion ....................................................................... 39
1.8 Diabetes ......................................................................................................................... 40
1.8.1 Type 1 diabetes ...................................................................................................... 41
1.8.2 Type 2 diabetes ...................................................................................................... 41
1.8.3 Monogenic diabetes ............................................................................................... 42
1.9 Genetics of T2D ............................................................................................................. 42
1.9.1 Single nucleotide polymorphisms (SNPs) ............................................................... 43
1.9.2 Genome wide association studies (GWAS) ............................................................ 44
1.9.3 SLC30A8 and T2D risk ............................................................................................. 45
1.10 The α-cell and glucagon secretion in T2D ................................................................... 47
1.11 Zn2+ in diabetes ............................................................................................................ 48
1.12 Cellular Zn2+ homeostasis ............................................................................................ 49
1.12.1 Metallothioneins .................................................................................................. 49
1.12.2 ZIP transporters .................................................................................................... 50
1.12.3 ZnT transporters ................................................................................................... 50
1.13 ZnT8 ............................................................................................................................. 51
1.13.1 ZnT8 and the β-cell; studies using mouse knockout lines .................................... 54
1.13.2 ZnT8 and the α-cell so far ..................................................................................... 56
1.14 Targeting glucagon for T2D treatment ........................................................................ 56
Aims of the thesis ................................................................................................................ 57
Chapter 2 – Materials and Methods ....................................................................................... 58
2.1 In vivo animal work ........................................................................................................ 59
2.1.1 Mice maintenance and generation ........................................................................ 59
2.1.2 Generation of transgenic mice ............................................................................... 59
2.1.3 Glucose tolerance tests .......................................................................................... 59
2.1.5 Insulin tolerance tests ............................................................................................ 59
2.1.6 Fasting plasma glucagon measurements ............................................................... 60
2.1.7 Streptozotocin treatments ..................................................................................... 60
2.1.8 Hypoglycaemic clamps ........................................................................................... 60
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2.2 In vitro experiments ...................................................................................................... 60
2.2.1 Islet preparation ..................................................................................................... 60
2.2.2 Islet dissociation ..................................................................................................... 61
2.2.3 Glucagon secretion ................................................................................................. 61
2.2.4 Ca2+ imaging with Fluo-2......................................................................................... 62
2.2.5 Zn2+ imaging with FluoZin-3 .................................................................................... 62
2.2.6 Granular Zn2+ imaging with Zinpyr-4 ...................................................................... 63
2.2.7 Mammalian cell culture .......................................................................................... 63
2.2.8 Zn2+ imaging with eCALWY-4: cells ......................................................................... 64
2.4.4 Amplification and purification of adenoviral eCALWY4 ......................................... 65
2.4.5 Virus titration.......................................................................................................... 65
2.2.9 Immunostaining: cells ............................................................................................. 66
2.2.10 Immunostaining: islets ......................................................................................... 66
2.2.11 Immunostaining: slices ......................................................................................... 67
2.2.12 Quantification of pancreatic glucagon content .................................................... 67
2.2.13 Electron microscopy ............................................................................................. 68
2.3 Molecular biology .......................................................................................................... 68
2.3.1 Genotyping ............................................................................................................. 68
2.3.2 RNA extraction ........................................................................................................ 68
2.3.3 cDNA conversion .................................................................................................... 69
2.3.4 Real Time PCR ......................................................................................................... 69
2.3.5 Fluorescence activated cell sorting (FACS) ............................................................. 69
2.3.6 FACS analysis .......................................................................................................... 70
2.4 Cloning and virus production ........................................................................................ 70
2.4.1 Competent bacteria ................................................................................................ 70
2.4.2 Transfection ............................................................................................................ 71
2.4.3 Bacterial transformation ........................................................................................ 71
2.5 Statistical analysis .......................................................................................................... 71
2.6 Primers .......................................................................................................................... 73
2.7 Antibodies ...................................................................................................................... 75
Chapter 3 - ZnT8 deletion from a subpopulation of α-cells enhances hypoglycaemia-induced
glucagon secretion .................................................................................................................. 76
3.1 Introduction ................................................................................................................... 77
3.2 Aims ............................................................................................................................... 78
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3.3 Results ........................................................................................................................... 79
3.3.1 Expression of ZnT8 in the mouse α-cell .................................................................. 79
3.3.2 Generation, breeding and phenotyping of mice null for ZnT8 in the α-cell ........... 80
3.3.3 Recombination efficiency and ZnT8 deletion ......................................................... 82
3.3.4 Body weight and growth ........................................................................................ 84
3.3.5 Glucose homeostasis .............................................................................................. 86
3.3.6 Insulin sensitivity .................................................................................................... 88
3.3.7 Streptozotocin treatment ....................................................................................... 89
3.3.8 Hypoglycaemic clamps ........................................................................................... 89
3.4 Discussion ...................................................................................................................... 91
3.4.1 Recombination in the α-cell ................................................................................... 91
3.4.2 Glucose homeostasis and insulin sensitivity .......................................................... 92
3.4.3 Response to hypoglycaemia in vivo ........................................................................ 93
3.5 Conclusion ..................................................................................................................... 93
Chapter 4 - ZnT8 is necessary in the α-cell for normal cytosolic and granular Zn2+ homeostasis
................................................................................................................................................. 94
4.1 Introduction ................................................................................................................... 95
4.2 Aims ............................................................................................................................... 97
4.3 Results ........................................................................................................................... 98
4.3.1 Impact of α-cell ZnT8 deletion on glucagon secretion in vitro ............................... 98
4.3.2 Calcium responses .................................................................................................. 98
4.3.3 α-cell area ............................................................................................................. 100
4.3.4 Intracellular Zn2+ imaging ..................................................................................... 100
4.3.5 Impact of α-cell ZnT8 deletion on secretory granule Zn2+ content ...................... 105
4.3.6 Gene expression analysis ...................................................................................... 105
4.3.7 Glucagon content ................................................................................................. 107
4.4 Discussion .................................................................................................................... 109
4.4.1 Glucagon secretion ............................................................................................... 109
Zn2+ homeostasis in the α-cell ....................................................................................... 109
4.4.2 Absence of effects of ZnT8 inactivation in α-cells on glucose-regulated Ca2+ fluxes
....................................................................................................................................... 110
4.5 Conclusion ................................................................................................................... 111
Chapter 5 - Overexpression of human ZnT8 in the murine pancreatic α-cell leads to reduced
glucagon release and impaired responses to hypoglycaemia ............................................. 112
5.1 Introduction ................................................................................................................. 113
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5.2 Aims ............................................................................................................................. 114
5.3 Results ......................................................................................................................... 115
5.3.1 Generation and genotyping of αZnT8Tg mice ...................................................... 115
5.3.2 Expression of human ZnT8 in the islet ................................................................. 116
5.3.3 Intraperitoneal glucose tolerance was unaltered in α-ZnT8Tg mice ................... 117
5.3.4 Intraperitoneal insulin tolerance tests (ITTs) ....................................................... 118
5.3.5 Hypoglycaemic clamps ......................................................................................... 118
5.3.6 Glucagon secretion is impaired in islets isolated from αZnT8Tg mice ................. 119
5.4 Discussion .................................................................................................................... 122
5.5 Conclusion ................................................................................................................... 123
Chapter 6 – Final Discussion .................................................................................................. 124
6.1 Investigating the role of ZnT8 in the pancreatic α-cell through mouse molecular
genetics ............................................................................................................................. 125
6.2 ZnT8 in the pancreatic α-cell ....................................................................................... 126
6.3 ZnT8 and Zn2+ homeostasis in the α-cell ..................................................................... 131
6.4 ZnT8, SLC30A8 and T2D risk ........................................................................................ 132
6.5 Final conclusions .......................................................................................................... 136
6.6 Future directions ......................................................................................................... 137
Chapter 7 - References .......................................................................................................... 138
Publications ........................................................................................................................... 156
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List of figures Figure 1.1 – Mouse pancreas, islets, α- and β-cells ................................................................. 18
Figure 1.2 – A simplified model for the role of transcription factors in endocrine
compartment differentiation .................................................................................................. 21
Figure 1.3 – Structure of proglucagon and processing............................................................ 23
Figure 1.4 Schematic model for KATP channel dependent glucagon secretion. ....................... 31
Figure 1.5 – Subcellular localization and direction of transport of the ZnT and ZIP Zn2+
transporter families. ................................................................................................................ 52
Figure 2.1 – eCALWY sensor design......................................................................................... 65
Figure 3.1 – ZnT8 expression in the α-cell ............................................................................... 79
Figure 3.2 – Generation of αZnT8 KO mice and genotyping by PCR ....................................... 81
Figure 3.3 – Breeding strategy ................................................................................................ 82
Figure 3.4 – Recombination in α-cells ..................................................................................... 83
Figure 3.5– ZnT8 deletion in α-cells. ....................................................................................... 83
Figure 3.6 – FACS analysis of WT and KO islets ....................................................................... 85
Figure 3.7 – Immunostaining of iGluCre islets ........................................................................ 86
Figure 3.8 – Weight measurements ........................................................................................ 87
Figure 3.9 – Glucose homeostasis ........................................................................................... 87
Figure 3.10 – Fasting glucose and glucagon ............................................................................ 88
Figure 3.11 – Insulin sensitivity ............................................................................................... 88
Figure 3.12 – Streptozotocin treatments ................................................................................ 89
Figure 3.13 – Hypoglycaemic hyperinsulinaemic clamps ........................................................ 90
Figure 4.1 - Glucagon secretion in vitro................................................................................... 98
Figure 4.2 – Low glucose-induced calcium responses. ............................................................ 99
Figure 4.3 – Pancreatic histology .......................................................................................... 100
Figure 4.4 – Zn2+ imaging with eCALWY-4 ............................................................................. 101
Figure 4.5 – Islet expression of adenoviral Zn2+ probe .......................................................... 102
Figure 4.6 – AAV-eCALWY-4 .................................................................................................. 103
Figure 4.7 – Intracellular Zn2+ imaging .................................................................................. 104
Figure 4.8 – Granular free Zn2+ imaging ................................................................................ 106
Figure 4.9 – Gene expression analysis ................................................................................... 107
Figure 4.10 – Glucagon content measurements ................................................................... 108
Figure 5.1 – Genotyping by PCR ............................................................................................ 115
Figure 5.2 – Human ZnT8 expression in isolated islets ......................................................... 116
Figure 5.3 – Hum an ZnT8 expression in purified α-cells ...................................................... 117
Figure 5.4 – Glucose tolerance in αZnT8Tg mice .................................................................. 117
Figure 5.5 – Insulin sensitivity ............................................................................................... 119
Figure 5.6 – Elevated glucose infusion rates in αZnT8 KO mice during hypoglycaemic
hyperinsulinaemic clamps ..................................................................................................... 120
Figure 5.7 - Glucagon secretion from αZnT8Tg islets is impaired in vitro ............................. 120
Figure 6.1 – Hypothetical model reconciling data on the effect of Zn2+ on glucagon secretion
and our mouse models .......................................................................................................... 129
Figure 6.2 – Hypothetical model reconciling data for rare and common SLC30A8 variants in
the β-cell ................................................................................................................................ 134
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Abbreviations
Arg arginine
Arx Aristaless Related Homeobox
BSA bovine serum albumin
DIO diet induced obesity
DMEM Dulbecco’s modified Eagle’s medium
FBP1 fructose-1,6-bisphosphatase
FRET fluorescence resonance energy transfer
F(1,6)P2 fructose-2,6-bisphosphate
GABA γ-aminobutyric acid
cAMP cyclic AMP
CcgR glucagon receptor
DMSO dimethyl sulfoxide
FACS fluorescence activated cell sorting
FBS fetal bovine serum
FRET förster resonance energy transfer
GIP gastric inhibitory polypeptide
GRPP glicentin-related polypeptide
GLP-1 glucagon-like peptide 1
GRPP glicentin-related polypeptide
GWAS genome wide association studies
G6PC glucose-6-phosphatase
HTRF homogeneous time resolved fluorescence
HVA high voltage activated
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IPGTT intraperitoneal glucose tolerance test
ITT insulin tolerance test
KATP ATP-sensitive K+ channel
LB luria broth
LepR leptin receptor
MOI multiplicity of infection
MT metallothionein
Ngn3 Neurogenin 3
Pax-4 Paired box-4
PBS phosphate buffered saline
Pdx-1 pancreatic and duodenal homeobox 1
PCR polymerase chain reaction
PCK2 phosphoenolpyruvate carboxykinase
Pcsk2 Proprotein Convertase Subtilisin/Kexin Type 2
PFA paraformaldehyde
PI3K phosphatidylinositol 3-kinase
PKA Protein Kinase A
PPG preproglucagon
RFP red fluorescent protein
RRP readily releasable pool
tTA tetracycline controlled trancactivator
rtTA reverse tetracycline controlled transactivator
SNP single nucleotide polymorphism
Tet tetracycline
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TPEN N,N,N′,N′-tetrakis(2-pyridylmethyl) ethylenediamine
Trp tryptophan
T1D type I diabetes
T2D type II diabetes
VIP vasoactive intestinal neuropeptide
VMH ventromedial hypothalamus
ZIP Zrt-, Irt proteins
ZnT zinc transporter
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Chapter 1
Introduction
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1.1 The pancreas
The pancreas is an essential organ in nutrient metabolism and serves as both an
exocrine and endocrine gland [1]. The exocrine pancreas, composed chiefly of acinar
cells, is involved in nutrient digestion through the production and secretion of
digestive enzymes into the intestine through the pancreatic duct. These include
enzymes such as carboxypeptidase-A, trypsin and amylase [2]. The endocrine
pancreas functions largely independently from the exocrine compartment and is
responsible for the maintenance of glucose homeostasis by secreting hormones into
the bloodstream. Endocrine cells cluster together to form the islets of Langerhans,
micro-organs that can be found scattered throughout the pancreas between the
exocrine tissue (Figure 1.1A) [3].
During embryogenesis, the pancreas is formed by fusion of a dorsal and ventral bud
created by evagination of the foregut. In the mouse the dorsal bud develops on
embryonic day 9.5 at the same time as the first differentiated glucagon expressing α-
cells develop [4]. Insulin-positive cells appear the next day, often simultaneously
producing glucagon [5]. By day 14, much larger numbers of β-cells can be seen and
the endocrine cells begin to arrange into small clusters. By the following day δ-cells
expressing somatostatin are present. Then, briefly before birth, the pancreatic
polypeptide (PP)-producing cells develop and the endocrine cells initiate the
formation of the highly organized islets of Langerhans [6].
1.2 Islets of Langerhans
1.2.1 Islet architecture
The islets of Langerhans account for 1-2% of total pancreatic mass only (~2000 in
rodents and 1 million per pancreas in humans). They tend to be spherical, 50-200 μm
in diameter and range in size from 1000 to 10000 cells [7]. There are several
different endocrine cell types in islets which work co-operatively to regulate glucose
metabolism: α-cells produce glucagon, β-cells insulin, δ-cells somatostatin, PP cells
pancreatic polypeptide. Additionally, a separate population of islet cells has been
identified the ε-cells which produce ghrelin, a hormone secreted mainly by the
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stomach to influence feeding behaviour [8]. The two main hormones are insulin and
glucagon which are released reciprocally in response to glucose fluctuations to keep
its levels within a relatively small range. Islets thus constitute a multihormonal
micro-organ, its sole purpose the maintenance of glucose homeostasis.
Rodent islets have a specific islet architecture where the vastly abundant β-cells are
clustered within the islet core surrounded by a thin layer of predominantly α-cells
along with the other endocrine cells (Figure 1.1B). This anatomical subdivision is not
present in human islets as the β-cells intermingle with α- and δ- cells throughout the
islet [9]. This lack of segregation of the different cell types in human islets has been
reported by various groups [9, 10] and could be suggestive of stronger paracrine
interactions compared to rodent islets. The cellular composition of islets also differs
between rodents and humans. In mouse islets [9] the β- cells make up the vast
majority of the islet reaching 70-80%, α-cells 15-20%, δ-cells 5-10% and the PP cells
<1%. In human islets the proportion of β-cells is only 50-60%, α-cells 30-40% and δ-
cells ~10% [9]. α- and β-cells are virtually indistinguishable from each other except
when achieving the high resolution of an electron microscope where the subtle
differences in the appearance of their secretory granules can help identify them
(Figure 1.1C,D). Imaged by conventional transmission electron microscopy β-cell
granules have a halo surrounding their dense core while α-cell granules appear
dense throughout [11].
1.2.2 Islet microvasculature
Pancreatic islets are highly vascularised with a dense network of sinusoidal
capillaries [12]. These capillaries have about 10 times more fenestrations than those
in the exocrine pancreas [13] that allow direct exchanges between the blood and
islet cells. In the rat, islets receive 10-15% of total blood flowing to the pancreas [14].
This direct interface allows the islet to respond rapidly to changing glucose
concentrations and other nutrients with hormonal release into the circulation. The
direction of blood flow within the islet is still an issue of debate. It has been
proposed that, in the rodent islet, blood flows initially through β-cell areas before
reaching non-β cell populations or that alternatively non-β-cells receive blood first
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before β-cells [15]. In human islets no anatomical subdivision was observed so that
all endocrine cell types were randomly distributed along blood vessels [9]. Such
findings suggest the theory that there is no specific direction of islet perfusion so
that each cell type could influence different cell types not excluding its own cell type
[15].
1.2.3 Islet innervation
Islets are extensively innervated to allow autonomic regulation over hormone
release. Both sympathetic (adrenergic) and parasympathetic (cholinergic) fibres
reach deeply into the islet. The sympathetic system is stimulated by hypoglycaemia,
exercise or stress, releasing noradrenaline [16] into the islet along with the
neuropeptides galanin [17] and neuropeptide Y [18]. Activation of the sympathetic
system triggers the secretion of glucagon with adrenaline and noradrenaline acting
via the α1- and β-adrenoceptors [19]. In contrast, adrenaline and the sympathetic
neuropeptides are inhibitors of both insulin [20] and somatostatin secretion [21].
The parasympathetic system which is activated during hyperglycaemic conditions
releases the neurotransmitter acetylcholine [22] and neuropeptides such as
vasoactive intestinal polypeptide [23]. Parasympathetic activation exerts an overall
stimulatory effect on the islet inducing secretion of insulin, glucagon, somatostatin
and PP [24, 25]. The majority of research on islet innervation has been focused on
the sympathetic system while the contribution of the parasympathetic mediators to
enhancing glucagon secretion is still unclear. Sensory neurons releasing γ-
aminobutyric acid (GABA) [26] and other neurotransmitters such as nitric oxide [27],
cholecystokinin [28], ATP [29], glutamate [30], dopamine and serotonin [31] have
also been identified in islets.
In humans, however, cholinergic innervation of islets is sparse [32]. In contrast to
mouse islets, acetylcholine acts as a paracrine signal released from α-cells affecting
the surrounding endocrine cells of the islet rather than being primarily a neural
signal.
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Figure 1.1 – Mouse pancreas, islets, α- and β-cells. (A) Representative image of a mouse
pancreas acquired by widefield microscopy. The green and red dots represent islets while
the rest is the exocrine pancreas. (B) Representative image of an islet stained for insulin
(green) and glucagon (red). Scale bar 50 µm. Transmission electron microscopy image of a
mouse α- cell (C) and β-cell (D). Note the difference in the structure of the dense core
vesicles between the two cell types. Scale bar 1 µm.
A
C B
D
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1.3 Glucagon
1.3.1 Historical perspectives
The history of glucagon started at the same time as that of insulin. When Banting
and Best tested their primary pancreatic extracts on pancreatectomised dogs, in
1921, they noticed that hypoglycaemia caused by insulin was followed by a mild
hyperglycaemia which they believed was a consequence of adrenaline release [6].
Murlin et al [33] discovered that injecting these pancreatic extracts caused an
increase in blood glucose in both healthy and diabetic dogs. They attributed this
effect to a glucogenic substance, and this was called glucagon [33]. Then in 1948
Sutherland and de Duve demonstrated, through a series of experiments, that the α-
cells of the pancreas were supplying the glucagon [34]. A few years later Foa et al
performed cross-circulation experiments in anaesthetised dogs which showed that
glucagon release is stimulated by hypoglycaemia [35]. Advances in glucagon
purification [36], and identification of its amino acid composition [37] subsequently
made studies into its physiology possible. In 1961, Unger et al developed glucagon
antibodies and a glucagon radio-immuno assay for assessing glucagon levels [38] and
in 1962 glucagon was localised to α- cells by immunofluorescence staining [39].
1.3.2 Pancreatic α-cell development
Multiple transcription factors have been discovered with temporally and spatially
limited expression during pancreas formation (Figure 1.2). During pancreatic
development, pancreatic duodenum homeobox-1 (Pdx-1) is expressed to promote
the development of the gland [40]. The transcription factor Neurogenin 3 (Ngn3), is
necessary to stimulate the next step, i.e. for progenitor cells to follow an endocrine
path. Ngn3 deletion in mice leads to an absence of pancreatic endocrine cells [41].
Neuro D is then needed for proliferation and is expressed in all endocrine cells [42].
Aristaless Related Homeobox (Arx) is responsible for correct endocrine development
as Arx-deleted mice exhibit an early onset loss of mature α-cells and at the same
time experience elevated numbers of β- and δ-cells [43]. Therefore, Arx is essential
for α-cell fate and to restrict β- and δ-cells. This function in endocrine commitment is
20
opposite to that of Paired box-4 (Pax-4) since there is a build-up of Pax-4 and Arx
transcripts when Arx and Pax-4 are deleted in mice, respectively [43]. This
antagonism between the two transcription factors for islet cell differentiation
determines the earliest stage of progression into α- or β-cells and is dependent on
the levels of each transcript [6].
Brain specific homeobox-4 (Brn-4) activates the glucagon gene [44] alongside MafB
which is solely expressed in the α-cell and further promotes cell type specific
transcription of glucagon [45]. Pax-6 is also necessary for differentiation into α-cells
determined by studies on mice which, when lacking Pax6, developed either very few
or no glucagon-expressing cells [46].
1.3.3 Preproglucagon – transcriptional control and processing
The proglucagon gene is expressed in pancreatic α-cells, intestinal L- cells, and in the
brain, primarily the hypothalamus and thalamus, hindbrain and olfactory bulb [6].
The preproglucagon gene was discovered and isolated from humans and rodents in
the 1980s. Following molecular cloning the isolation of hamster preproglucagon
cDNA [47] and the human glucagon gene [48] revealed the existence of three
homologous sequences, glucagon, GLP-1 and GLP-2 [48, 49].
These are joined together by two intervening peptides IP-1 and IP-2 and are
downstream of the signal peptide glicentin-related polypeptide (GRPP) at the N
terminus.
Mammalian preproglucagon is composed of 179-180 amino acids. The three
peptides are bound to each other by a pair of basic amino acids, potential sites for
proteolytic cleavage by proteases involved in hormone maturation [50].
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Figure 1.2 – A simplified model for the role of transcription factors in endocrine
compartment differentiation. The position of each transcription factor indicates its time of
expression or its main functional role. The most important transcription factors in the
differentiation of α- and β-cells are shown. Some are expressed at several steps but they are
only depicted once for simplicity. From Gromada et al [6].
22
The hormone that will eventually be produced from preproglucagon depends on the
presence of specific proteases known as prohormone convertases. Proprotein
Convertase Subtilisin/Kexin Type 2 (Pcsk2) is exclusively found in the α-cell and is
responsible for processing proglucagon into active 29 amino acid-long glucagon,
GRPP and IP-1, peptides with unknown functions, and the major proglucagon
fragment (MPGF) of inactive GLP-1, IP-2 and GLP-2. Conversely, in the L cell the
presence of Pcsk1/3 and the lack of Pcsk2 lead to the production of GLP-1 and GLP-2
alongside IP-2 and glicentin, the N-terminal fragment that includes unprocessed
glucagon which is processed to GRPP and oxyntomodulin (Figure 1.3).
The promoter of the preproglucagon gene has six key elements upstream the
transcription initiation site. These sequences of 20-40 bp are called G1, G2, G3, G4,
CRE (cAMP response element) and ISE (intestinal specific element). The G1 sequence
allows α-cell specific expression of the gene [51] while G2-G4 are islet specific
enhancers [52]. The CRE sequence is responsible for transcription in response to
cAMP [53] and ISE confers expression specifically in the L-cell [54]. Pax-6 is the main
transcription factor during pancreas development through transactivation of G1 [55],
while Pax-4 inhibits glucagon expression by preventing Pax-6 transactivation [56].
Pdx-1 acts as a repressor of glucagon transcription [57] and is important for islet cell
differentiation being an important transactivator of the preproinsulin gene [58].
Insulin has been shown to act as an inhibitor of glucagon expression in α-cells by
affecting transcription rates through an insulin-responsive DNA element in the
proximal 5’ flanking region of the G3 promoter sequence [59]. Later studies proved
that in addition to the proximal DNA regions distal promoter elements are involved
in the insulin mediated inhibition of glucagon expression [60].
23
Figure 1.3 – Structure of proglucagon and processing. Top: structure of proglucagon. At the
end of each peptide there are pairs of dibasic amino acids (KK, RR, KK where K is lysine, R
arginine) and these are potential cleavage sites by prohormone convertases. Bottom left:
Proglucagon processing occurring in the α-cell. The peptides are GRPP, glucagon, IP1 and
MPGF which can be partially processed to GLP-1. Bottom right: Processing of proglucagon in
the L cell. The products are glicentin, truncated GLP-1, IP2 and GLP-2. Glicentin can be
partially processed to GRPP and oxyntomodulin. Adapted with permission from Furuta M et
al., J. Biol. Chem., 2001 [61].
GRPP
glucagon
IP1
GRPP glucagon GLP-1 GLP-2
KR KR KR RR RR KK
COOH
IP1 IP2
KR
GLP-1
MPGF
PC2 PC1/3
GRPP
GLP-1
IP2
Oxyntomodulin
GLP-2
Proglucagon
α-cell L cell
Glicentin
24
1.3.4 Glucagon receptor
The rodent glucagon receptor (GCGR), composed of 485 amino acids is a member of
the secretin-glucagon receptor class II family and is G protein-coupled receptor [62].
The binding of glucagon is coupled to a Gαs type GTP-binding G protein that in turn
causes the activation of adenylate cyclase and subsequently production of cAMP and
Protein Kinase A (PKA) activation. Glucagon binding can also induce the
phospholipase C/inositol pathway through Gq proteins, mobilising intracellular Ca2+
stores [62]. The glucagon receptor is found in several tissues being most abundant in
the liver and kidney and to a lesser extent in the endocrine pancreas, heart, cerebral
cortex, adipose tissue and smooth muscle regulating various processes in these
tissues to modulate glucose homeostasis [63].
Genetic polymorphisms in the GcgR have been found to be linked to Type 2 diabetes
(T2D). For example a missense mutation in exon 2 of the GcgR gene that causes an
amino acid change is associated with T2D in French populations. The mutated
receptor appears to have a decreased affinity for glucagon and lowered cAMP
responses after stimulation [64]. However, the mutation was not associated with
T2D in other populations [65]. Glucagon receptor null mice have lower glucose levels
although they are not hypoglycaemic but display α-cell hypertrophy and severe
hyperglucagonaemia [66].
1.3.5 Physiological actions of glucagon
The body has developed several ways to defend against hypoglycaemia and the likely
harmful effects that could ensue, particularly in the brain, which relies heavily on a
steady supply of glucose as a fuel. Such defences include a reduction in insulin
secretion and elevated release of adrenaline and glucagon [67]. Glucagon has a key
role in the response to hypoglycaemia, its primary action being in the liver where the
ratio of insulin/glucagon regulates various aspects of hepatic metabolism. Thus,
glucagon induces gluconeogenesis and glycogenolysis resulting in an increase of
hepatic glucose production for release into the bloodstream and subsequent supply
of glucose to the rest of the body and very importantly to the brain [68]. It
25
simultaneously decreases glycolysis and glycogenesis which would use up glucose.
Through the highly selective glucagon receptor, glucagon mediates its actions in the
liver mainly by activating the adenylyl cyclase and the PKA pathway. Through this
mechanism, glucagon elevates gluconeogenesis by the upregulation of enzymes
including glucose-6-phosphatase (G6PC) and phosphoenolpyruvate carboxykinase
(PCK2) via induction of the cAMP response element-binding protein (CREB) and
Peroxisome proliferator-activated receptor gamma 1 (PPARγ1) [69, 70]. Another
important player is coactivator transducer of regulated CREB activity 2 (TORC2).
Glucagon causes the dephosphorylation and subsequent translocation of TORC2 to
the nucleus where it enhances CREB-dependent transcription [71]. PCK2, G6PC and
fructose-1,6-bisphosphatase (FBP1) are important in the rate of gluconeogenesis.
PCK2 is responsible for the conversion of oxaloacetate into phosphoenolpyruvate
and G6PC catalyses glucose transition of glucose-6-phosphate to glucose. FBP1
mediates production of fructose-6-phosphate from fructose-1,6-bisphosphate
(F(1,6)P2). The activity of this enzyme is controlled by glucagon as it reduces the
levels of fructose-2,6-bisphosphate (F(2,6)P2), its allosteric inhibitor [72]. This
reduction of F(2,6)P2 in turn reduces phosphofructokinase-1 activity, decreasing the
flux through glycolysis. Glycolysis is also inhibited by glucagon at the pyruvate kinase
step [73].
The direction of glycogen metabolism is determined primarily by the action of
glycogen synthase (GS) and glycogen phosphorylase (GP). Glucagon mediates GP
phosphorylation and activation and at the same time inactivates GS by
phosphorylation thus causing glycogenolysis [74, 75]. Additionally, in aid of elevating
glucose levels glucagon stimulates the uptake of amino acids by the liver for
gluconeogenesis [76].
1.4 Glucagon secretion
Each α-cell contains ~ 7500 secretory granules [77]. These granules have a similar
sized dense core to β-cells with a core diameter of ~230 nm. They are round and
have a dark electrondense core but lack the characteristic halo of insulin containing
26
granules making them smaller in overall diameter [78]. About 1% of these granules
are docked beneath the plasma membrane and make up the readily releasable pool
(RRP) [77].
1.4.1 Ion channels in the α-cell
α-cells differ sharply from β-cells in their response to glucose, generating action
potentials in the absence of sugar or at sub-physiological levels. Activation of
voltage-gated Na+ and Ca2+ currents is needed for the initiation of the action
potential and K+ currents are necessary for spike repolarization. Several types of ion
channels are found in the α-cell. There are four types of K+ selective channels; the
ATP sensitive K+ channel (KATP), the delayed rectifying K+ channel, a G-protein gated
K+ channel stimulated by somatostatin and a transient K+ channel. Also identified are
four types of voltage gated Ca2+ channels (T, N, R, L), a Na+ channel and GABA
receptor chloride channels [6].
KATP channels have a key function in coupling cell metabolism to electrical activity in
a variety of tissues [79]. In the β-cell, during resting conditions, the basal activity of
the channel sustains the negative membrane potential until glucose metabolism
leads to a concentration-dependent inhibition of the channel, due to a change in the
ATP/ADP ratio within the cell. KATP closure causes membrane depolarization and the
subsequent opening of voltage-dependent Ca2+ channels. Ca2+ influx then ensues
followed by Ca2+ dependent exocytosis [6]. The KATP channel in α-cells is highly
sensitive to ATP and thus more sensitive to inhibition than in the β-cell requiring
concentrations as low as 0.05 mM to cause maximal stimulation of activity[6].
Voltage-dependent K+ channels are necessary for repolarization of the α-cell action
potential, while Na+ channels are required for the production of an action potential.
Delayed rectifying K+ currents have been detected in mouse, guinea pig and in rat α-
cells [77, 80]. This current is sensitive to tetraethylammonium which can decrease
the amplitude of the outward current by >90%. Its application to rat α-cells led to
enhanced glucagon secretion due to elevated membrane depolarisation [81].
27
The Na+ channels are tetrodotoxin-sensitive and are functional at potentials ranging
from positive to -30 mV in mouse α-cells. Their function is essential for generation of
action potentials. Tetrodotoxin suppresses glucagon secretion from rodents to a
level similar to that seen by high glucose [82].
The voltage-gated Ca2+ channels the α-cell is equipped with belong to three
subfamilies which are: 1) the L-type high voltage-activated (HVA) channels, 2) P/Q-,
N-, and R- type HVA Ca2+ channels 3) the low voltage activated T-type channels [80].
The opening of the T-type channels takes place at membrane potential as low as -65
mV while for HVA channel activity depolarisations of -40 to -30 mV and above are
needed. The different requirements for voltage values indicate that the low and high
voltage channels could serve separate functions in action potential generation. T-
type channels are active around the action potential threshold and it has been
suggested that they are involved in action potential initiation. HVA channels open at
membrane potentials near the rising phase of the action potential and are of greater
capacity accounting for the majority of the Ca2+ entry necessary for glucagon release
[6].
1.4.2 Regulation by glucose: paracrine or intrinsic?
The question of whether the inhibition of glucagon secretion by glucose occurs due
to the release of factors from neighbouring cells or whether α-cells have the intrinsic
ability to sense changes in glucose concentrations is a matter of debate. Insulin and
glucagon secretion being reciprocally regulated by glucose prompted the speculation
that glucagon secretion is controlled in a paracrine manner by insulin which is known
as the ‘switch-off hypothesis’ [83]. Such a hypothesis would easily explain the
hypersecretion of glucagon observed in diabetes, it being a result of low insulin
release, and why this is abolished with the aid of exogenous insulin. Variations to the
concept propose factors released with insulin from secretory granules could be
contributing to or be responsible for the suppressive effect of high glucose on α-cells
[84].
28
The issue with this hypothesis is that the α-cells are maximally inhibited at glucose
levels below the threshold for β-cell activation and insulin secretion or somatostatin
[85]. Studies indicate that isolated single α-cells still respond to glucose by lowering
intracellular Ca2+ concentrations and suppression of glucagon release [86, 87]. The
suppressive effect of glucose was preserved when insulin signalling was blocked
using wortmannin [88]. Additionally there is evidence of vagal control of glucagon
secretion [89], yet following denervation the α-cell response to hypoglycaemia is
intact [90]. Such findings suggest that although the α-cells might be under paracrine
regulation they also possess an intrinsic ability to sense glucose.
This mechanism remains poorly defined but research on mice lacking functional KATP
channels, which have an impaired regulation of glucagon secretion by glucose,
suggests that KATP channels are involved in some way [91]. Mice lacking the KATP
channel subunit Sur1, in hypoglycaemic conditions, release similar amounts of
glucagon to WT islets exposed to maximally inhibitory glucose concentrations and
glucagon secretion cannot be inhibited further when the glucose concentration is
increased. Additionally, in WT mouse islets, the suppression of glucagon release by
high glucose can be reversed by low concentrations of diazoxide, a KATP channel
activator [82]. Human α-cells similarly express KATP channels and administering the
KATP channel blocker tolbutamide results in depolarisation suggesting that these
channels are active in the α-cell and have a repolarizing role [92].
1.4.3 Kinetics of exocytosis
Exocytosis of glucagon from the α-cells is Ca2+ dependent just like secretion of insulin
from β-cells, but is slightly faster. The difference is a result of the quantity of
granules in the RRP, approximately 120 in the α-cell compared to ~60 in the β-cell,
and the rate at which the RRP is refilled, ~ 20 granules/sec in the α-cell and 5-10
granules/sec in the β-cell [93]. Additionally, there is a robust secondary acceleration
of the exocytotic rate in the α-cell following the emptying of the RRP which could be
due to fast Ca2+ dependent translocation of secretory granules deeper into the cell to
29
the release sites. The maximum exocytosis rate of 750 granules/sec during
depolarisation is completed within 20 msec and is similar to that observed in β-cells
which is ~700 granules/sec and finishes in 15 msec [93].
Considering such findings it appears that the two cell types share a comparable
organization of Ca2+-dependent exocytosis, but suggest a higher number of release
sites is present in α-cells. The large Ca2+ currents of the smaller α-cells, resulting in
larger and less localised Ca2+ transients, could at least partly account for the
differences between the two cell types. Such features allow the α-cell to act rapidly
and secrete glucagon during short periods where glucose is in demand such in cases
of intense physical activity [93].
1.4.4 Stimulation of exocytosis
α-cells are electrically excitable and use electrical signals to couple any changes in
glucose levels to changes in glucagon output. They fire action potentials in the
absence of glucose, possessing a specific set of channels to generate currents of Na+
and Ca2+, and this electrical activity in turn produces Ca2+ signals and glucagon
release.
A key role for KATP channels is the widely most accepted hypothesis for glucose
sensing and glucagon secretion (Figure 1.4). In the mouse, at low glucose
concentrations, the function of the KATP channels is to keep the membrane potential
at ~-60 mV which is the threshold for action potential initiation. At such a voltage, T-
type Ca2+ channels become active bringing the membrane potential within a range
that allows activation of Na+ and HVA L-type and N-type Ca2+ channels which open at
membrane potentials more positive than -30mV. This leads to the regenerative
opening of these channels and action potential generation [85]. The entry of Ca2+
through N-type channels is what stimulates glucagon release. Repolarisation of
action potentials is achieved by the efflux of K+ through voltage dependent K+
channels.
30
An increase in glucose leads to a rise in cytosolic ATP/ADP levels which causes the
closure of the KATP channel. Subsequently the α-cell is depolarised but, unlike in the
β-cell the depolarisation suppresses glucagon release. The α-cell action potential
depends on T-type Ca2+ channels and Na+ channels and at the membrane potential
following sustained depolarisation the channels become inactivated thus inhibiting
electrical activity, Ca2+ fluxes and glucagon release [85, 94]. Therefore glucagon
secretion is primarily supported by an intermediate KATP channel activity which keeps
the membrane potential within a range that can promote regenerative action
potentials [85]. A similar mechanism has been suggested for glucagon secretion by
human islets [85].
The above mechanism has been questioned since it has been reported that glucose
may have a hyperpolarising instead of a depolarising effect [95, 96]. An alternative
model has been proposed of glucose suppressing glucagon release independently of
KATP channels by inhibition of a depolarizing Ca2+ store-operated current [95, 97].
1.5 Metabolism of the α-cell
1.5.1 Glucose
It has been well established that glucose triggers monophasic suppression of
glucagon secretion from islets. Based on dose-response experiments the α-cell
appears to be more sensitive to glucose compared to the β-cell [98]. The threshold
for glucagon inhibition is 2-3 mM glucose which is lower than the β-cell threshold of
4-5 mM which acts as a stimulus for insulin release. Additionally half-maximal
inhibition of glucagon is at 3-6 mM, while half-maximal stimulation of insulin
happens at 8-10 mM. Maximal suppression of glucagon secretion is achieved at 6-10
mM glucose, but for maximal stimulation of insulin release the glucose
concentration has to be above 20 mM [98, 99]. This difference in sensitivity makes it
unlikely that increased insulin secretion as glucose concentrations rise is responsible
for the inhibition of glucagon release through a paracrine mechanism. Of note, other
31
Figure 1.4 Schematic model for KATP channel dependent glucagon secretion. Under low
glucose conditions (top) the ATP/ADP ratio is low so KATP channels still maintain a low level of
activity. This causes moderate depolarisation which activates Na+ channels resulting in
action potential generation and a Ca2+ flux through Ca2+ channels. This leads to Ca2+-
dependent exocytosis of glucagon. At high glucose levels (bottom) there is increased entry of
glucose through Glut2 transporters which increases the ATP/ADP ratio and causes the
closure of KATP channels. This in turn inactivates Na+ channels so that there is no action
potential generation and no exocytosis. Adapted from Zhang et al [100].
glut 1
glut 1
32
related sugars and glucose metabolites have the same inhibitory impact on glucagon
secretion and the suppressive effect is based on the ability of the α-cell to
metabolize the sugar [101]. This observations is in agreement with the fact that
suppression of glucagon secretion by glucose is prevented by inhibitors of cellular
metabolism [102].
α-cells like β-cells express the high-Km glucokinase whose activity in the α-cell is
similar to that in the β-cell and is thought to have a glucose sensing function [103].
Additionally there is expression of the low-Km hexokinase II which is responsible for
~50% of glucose phosphorylation in α-cells compared to less than 10% in β-cells
[103]. This flux may be have a role in sensing very low glucose levels as it reaches
saturation at 1mM glucose [103].
The glucose transporter Glut1 is expressed in the α-cell instead of the more efficient
(but low affinity) Glut2 found in the rodent and human β-cells [104]. The lack of
Glut2 and the low levels of Glut1 mean that the uptake rate of glucose by the α-cells
is 10 times less than that by β-cells yet is it still 10 fold greater than the metabolic
rate so that transport is not a rate limiting factor in terms of glucose metabolism
[104]. The rate of glucose utilization i.e. glycolysis is similar in α-cells and β-cells
[104]. However glucose oxidation, i.e. oxidative phosphorylation, in α-cells reaches
only 10% of that in the β-cell [105]. Non quantitative, time-resolved measurements
of cytosolic ATP levels using a luminometric method have shown that glucose
increases induce a rapid increase in ATP [88]. Measuring ATP and ADP
concentrations biochemically in pure α-cells and β-cells indicate that the ATP/ADP
ratio in α-cells at 1mM glucose is the same as that in β-cells subjected to 6 mM
glucose [105].
1.5.2 Amino acids
Amino acids have a stimulatory effect on glucagon secretion, with an action on the
α-cell which is more potent than that on the β-cell [106]. The capacity of each amino
acid to stimulate glucagon secretion differs, with arginine being the most effective,
alanine and glutamine being potent but less so [86]. Some amino acids have little or
33
even no effect individually, but can potentiate the stimulation of glucagon secretion
by other amino acids [86]. Amino acids that overall have a small impact on insulin
secretion have a key role in the regulation of glucagon secretion under physiological
conditions. Since glucose enhances insulin responses to amino acids while
dampening glucagon release, amino acids in combination with glucose play a crucial
part in the differential stimulation of the cell types. Insulin release after a protein
meal would lead to hypoglycaemia if it was not accompanied by a stimulation of
glucagon secretion [6].
1.5.3 Fatty acids
Fatty acids influence glucagon release but the extent varies among different species
and experimental conditions [107]. The initial observation was that they have a
suppressive effect on glucagon release [108], but later studies showed that short
term exposure actually induces its secretion [109]. The short term stimulatory effect
of fatty acids on mouse islets is depended on chain length, the longer the chain the
higher the glucagon release. Spatial configuration is also important, with trans fatty
acids being more potent than cis isomers. The level of saturation similarly impacted
efficacy with saturated fatty acids being more effective than unsaturated [110].
Palmitate can increase exocytosis by enhancing Ca2+ influx through L-type Ca2+
channels and by reducing somatostatin release from δ-cells by 50% thus alleviating
the α-cell from its inhibition [111]. The long term effect of palmitate and oleate on α-
TC1.6 cells was to enhance glucagon secretion in a dose dependent manner but
suppress proliferation [112]. Consistent with these findings, long term exposure of
rat islets led to an enhancement in glucagon secretion, a reduction in glucagon
content likely a result of increased release since glucagon gene expression was
unaltered [113].
1.6 Regulators of glucagon secretion – autocrine, paracrine and endocrine
While the regulation of insulin secretion is well understood, the regulation of
glucagon secretion is a lot less clear with many hypotheses on how various factors
might affect it in a paracrine manner, being released from neighbouring cells, or how
34
it might be centrally controlled through innervation. These are discussed in more
detail below.
1.6.1 Insulin
β-cell secretory granules contain a high concentration of crystalline hexameric insulin
(~118 mM in human and 74 mM in rat primary cells [114]) and once released the
insoluble hexamers separate into the active monomeric form. It can thus be found at
a high local concentration in the islet with the potential to influence adjacent cells.
Insulin has long been regarded as the most likely contender of all the β-cell products
to have an inhibitory effect on secretion from α-cells [115]. In diabetic and non-
diabetic humans intravenous infusion of insulin, while maintaining normoglycaemia,
resulted in a reduction in glucagon secretion [116]. The effect of insulin on glucagon
secretion appears to be conserved across species. Thus exogenous administration of
insulin has been shown to inhibit glucagon release from perfused streptozotocin-
treated rat pancreas [117], streptozotocin-treated hamsters [118] and guinea pig
islets [119], alloxan-treated diabetic dogs [120], rat pancreas and isolated α-cells
including isolated rodent islets, but only at low glucose [121], and clonal mouse α-
cells [122]. Conversely, perfusion of isolated rat [123] and human [124] pancreata
with anti-insulin antibodies at 5.5 and 11.5 mM glucose respectively stimulated
glucagon release.
Insulin receptor transcripts are highly expressed in rat α-cells [121] and in clonal
mouse and hamster α-cells [122]. Insulin reduced glucagon secretion from both
mouse and hamster cell lines [122]. Deletion of the insulin receptor in mice
specifically in the α-cell caused mild glucose intolerance, hyperglycaemia and
hyperglucagonaemia in the fed state as well as an elevated response to arginine
[125]. Various groups have provided some insight into the mechanism by which
insulin acts in the α-cell. Evidence from rat α-cells suggests insulin (0.1 µM) has a
hyperpolarising effect on the α-cell by increasing KATP channel activity thus inhibiting
electrical activity and glucagon secretion [121]. In another study on primary mouse
α-cells, 3 µM insulin significantly reduced the sensitivity of KATP channels to inhibition
by ATP shown to be mediated via the phosphatidylinositol 3-kinase (PI3K) signalling
35
pathway [126]. Further to its effect on KATP channels, insulin, via the activation of a
PI3K-Akt dependent pathway, can cause the translocation of GABA-A receptors to the
plasma membrane leading to an increased response to GABA secreted by β-cells,
membrane hyperpolarization and suppression of glucagon release [127].
1.6.2 Zinc
In its crystalline form in secretory granules, insulin is centrally coordinated to two
Zn2+ ions to form hexamers [128]. Zn2+ levels in these granules reach ~20 mM [129]
and upon exocytosis the ions dissociate and are released into the microvasculature
or interstitial space [121] achieving a high local concentration with the potential to
act as mediators of paracrine signalling. An inhibitory effect of Zn2+ on glucagon
secretion was first observed when the Zn2+ chelator Ca2+-EDTA allowed stimulation
of secretion by the normally inhibitory mitochondrial substrate monomethyl-
succinate in the perfused rat pancreas while insulin secretion was unchanged [84]. In
support of this finding, Zn2+ was found to inhibit pyruvate- or glucose-induced
glucagon secretion from isolated rat α-cells by reversibly activating KATP channels and
thus reducing electrical activity and glucagon release [121]. Another study
demonstrated this inhibitory effect of Zn2+ treatment on glucagon secretion in both a
glucagon producing mouse cell line and intact mouse islets [130]. However, this was
not replicated in a similar study which showed no effect of Zn2+ on glucagon
secretion from mouse islets and a murine α-cell line [88].
1.6.3 Somatostatin
Somatostatin is widely distributed throughout the body being produced by both the
central and peripheral nervous system but also peripheral organs such as gut,
kidneys, adrenal and thyroid glands and the δ-cells of the pancreas [131].
Immunocytochemical studies revealed that out of the five somatostatin receptor
(SSTR) subtypes SSTR2 is highly expressed in α-cells while SSTR1 and SSTR5 are
expressed in β-cells in human islets [132]. In rodents SSTR2 and SSTR5 predominate
in the α-cells and β-cells respectively [133, 134]. Somatostatin has been established
36
as an inhibitor of both insulin and glucagon secretion [134, 135], released in the islet
is response to glucose with a threshold concentration of 4-5 mM.
Three mechanisms have so far been described as to how activation of the
somatostatin receptors causes the inhibition of glucagon release. Electrophysiology
data indicate that activation of G protein-coupled K+ channels by somatostatin leads
to hyperpolarisation of the cell membrane in isolated mouse and rat α-cells so that
electrical activity is inhibited [136, 137]. Somatostatin has also been found to
stimulate a Gαi2 protein-dependent pathway followed by localised activation of the
phosphatase calcineurin that in turn causes depriming of docked secretory granules
in primary rat cells. This effect did not involve a change in Ca2+ levels and
hyperpolarisation [138]. Additionally, somatostatin binding to its receptor inhibits
adenylyl cyclase activity and thus decreases cAMP levels and PKA induction of
glucagon release [139].
1.6.4 GABA
GABA is a major inhibitory neurotransmitter in the central nervous system but it is
also produced by β-cells which package it into microvesicles similar to synaptic
vesicles for release in a controlled manner [140]. GABA was shown to cause
hyperpolarization of rat α-cells upon activation of the GABAA receptor. The use of an
antagonist to explore the effect of endogenous GABA was found to increase
glucagon release from rat islets twofold at 1mM glucose and eliminated the
inhibition imposed by 20 mM glucose. However, the levels of secretion were still
lower than at low glucose conditions [141]. Exogenous application of GABA exerted
an inhibitory effect on arginine-induced glucagon release in guinea pig [142] and
mouse islets [143], primary rat α-cells [81] and the mouse line αTC6 [144].
1.6.5 Glutamate
Islet endocrine cells appear to have glutamatergic signalling elements with the ability
to make, release and respond to glutamate, the major excitatory neurotransmitter in
the CNS [145]. Glutamate is produced in β-cell mitochondria and in the cytosol, likely
37
accumulating in insulin granules with the possibility of acting as an intercellular
signalling agent following exocytosis [146]. Several of the subunits of the large
glutamate receptor family have been found to be expressed in the different types of
islet cells. In the α-cell both ionotropic receptors, AMPA and kainite subtypes, have
been identified [147] as well as metabotropic receptors [148]. The ionotropic
receptors act as Na+ channels whose activation regulates plasma membrane
potential and thus glucagon release. In neurons, the G-protein coupled metabotropic
receptors, modify cAMP levels and are involved in autocrine feedback inhibition of
glutamate signalling. Binding of glutamate to AMPA or kainite receptors in a
hypoglycaemic setting stimulated glucagon secretion from the perfused rat pancreas
[149]. Another study showed a modest inhibition of glucagon secretion from isolated
islets and the perfused rat pancreas following administration of GABA, an effect
believed to involve ionotropic receptors [143].
Vesicular glutamate transporter subtypes 1 and 2 have been identified in α-cell
glucagon granules and a concomitant elevation in glucagon and glutamate release
has been observed from rat islets in low glucose conditions [150]. Consequently,
glutamate co-released with glucagon might have a positive autocrine effect on
glucagon secretion. In contrast, the activation of α-cell metabotropic receptors leads
to suppression of glucagon release from rat islets under hypoglycaemic conditions
[151]. The evidently opposing effects of the two different types of glutamate
receptors following activation on glucagon release augment the level of complexity
regarding intra-islet glutamate signalling.
1.6.6 Glucagon-like-peptide (GLP-1)
GLP-1, an incretin secreted by L-cells in the small intestine following a meal [152],
primarily acts to enhance glucose-induced insulin release through GLP-1 receptors
on β-cells and via activation of the autonomic nervous system. GLP-1 also acts on the
α-cell to inhibit glucagon release although the expression of GLP-1 receptors in α-
cells is <0.2% of that in β-cells [153]. The suppressive ability of GLP-1 was observed in
vivo and in the perfused pancreas. One group concluded that its inhibitory effect
38
must be paracrine since in single isolated cells GLP-1 actually stimulated glucagon
release through interaction with G-protein couple receptors activating adenylate
cyclase which leads to cAMP production [154]. However, similar research on both
intact and isolated cells concluded that suppression of glucagon release by GLP-1 is
PKA-dependent, glucose-independent and does not involve paracrine mechanisms.
The incretin appears to inhibit not by affecting electrical activity rather by selective
inhibition of N-type Ca2+ channels and thus exocytosis [153].
1.6.7 ATP
ATP is another potential paracrine inhibitor of glucagon release. It can be found in
synaptic vesicles of islet nerve endings but also in insulin secretory granules [155].
Using immunocytochemistry α-cells were shown to express the purinergic receptors
including the P2 receptor subtype P2Y1 and the P1 receptors P1A1 and P1A2A [156].
Applying ATP extracellularly diminished or reduced the frequency of calcium
oscillations observed in α-cells at low glucose both in intact mouse islets and isolated
cells. This effect appears to be directly through ATP and not due to its hydrolysed
products and most likely involve P2Y1 and A1 receptors [156]. Previous findings from
perfused rat pancreata [157] and rat islets [158] contradict the inhibitory effect of
ATP on mouse islets since adenosine and ATP respectively stimulated glucagon
release. Such contrasting results could be due to species differences.
1.6.8 Glucagon
Glucagon has a stimulatory effect on insulin and somatostatin release from the
perfused pancreas of both rats and humans [124, 159]. It also has an autocrine role
having a positive feedback effect by binding to glucagon receptors expressed in α-
cells, elevating cAMP levels and exocytosis from both rat and mouse α-cells [160].
Experiments on mice with an inability to produce glucagon due to the absence of the
processing enzyme PCSK2 [161] or missing the glucagon receptor [162] suggest that
the lack of this autocrine control by glucagon or the loss of glucagon activity in target
tissues causes α-cell hyperplasia. Reducing glucagon receptor activity specifically in
the mouse liver led to enhanced glucagon secretion in vivo, but there was no
39
increase in α-cell mass indicating that the trigger stimulating hyperplasia was most
likely an autocrine effect and not due to the lowering of glucagon signalling in the
liver [163].
1.6.9 Ghrelin
Ghrelin is a growth hormone-releasing peptide synthesised by the stomach in fasting
conditions. It is also found in the human and rodent islets, released from a separate
cell population distinct from all other cell populations (α, β, δ, PP) of the islet [8]. It
appears that ghrelin-positive cells are more abundant in the foetal and neonatal
stage, diminishing in numbers at the adult stage, and seen mainly in the periphery of
the islet [164]. The ghrelin receptor is widely expressed in the islet and was observed
to colocalize with α-cells [165]. Ghrelin had a stimulatory effect on glucagon
secretion from isolated mouse islets, but no such effect was observed in vivo [166].
Additionally, and more recently ghrelin, was found to increase glucagon release from
mouse αTC1 and InRG9 guinea pig cell lines but also from isolated islets in a dose
dependent manner [167].
1.7 Neural regulation of glucagon secretion
As described earlier (Section 1.2.3) islets are amply innervated by sympathetic and
parasympathetic nerves to ensure rapid prevention or correction of hypoglycaemia
for protection of the brain [168]. Reductions in arterial glucose levels are sensed in
various areas of the brain, carotid bodies and the hepatic portal vein. A specific
region in the brain, the ventromedial hypothalamus (VMH) has been identified as an
important player in sensing falling blood glucose concentrations and also in
commencing counterregulatory measures against hypoglycaemia [169, 170]. There
are three levels of autonomic control over the α-cell, sympathetic and
parasympathetic nerves and circulating adrenaline, all of which are activated by
hypoglycaemia.
Stimulation of the sympathetic and parasympathetic arms of the nervous system is
achieved by glucose-sensing neurons in the VMH. The glucose sensing process is
40
thought to involve KATP channels since mice lacking a component of these channels
have a permanently raised firing rate and an extremely defective glucagon counter-
regulatory response [89]. Activating the channels in the VMH neurons
pharmacologically leads to enhanced adrenaline and glucagon responses during
hypoglycaemia [171].
Adrenaline is a strong stimulator of glucagon release in rodent islets caused by an
increase in cAMP levels, an effect which can be reproduced by using the adenylate
cyclase activator forskolin [153]. However, adrenaline appears to have no such
ability on glucagon release in human islets and similarly forskolin had no effect either
[172]. A stronger counterregulatory response by α-cells in response to adrenaline
could be a likely explanation as to why mice manage insulin-induced hypoglycaemia
better than humans [172].
Parasympathetic and sympathetic nerve terminals may contain stores of
neurotransmitter including acetylcholine and noradrenaline as well as several
neuropeptides. Once released these may either stimulate or suppress glucagon
depending on the specific peptide found in each terminal [63]. Cholinergic activation
through muscarinic receptors leads to cytoplasmic Ca2+ mobilization, increasing
glucagon release, as observed in isolated mouse α-cells and also in vivo [173]. The
same effect was detected after noradrenaline administration, which caused
adenylate cyclase activation and subsequently increased intracellular Ca2+
concentration and glucagon secretion [174]. Additionally, neuropeptides including
vasoactive intestinal neuropeptide (VIP) and gastric inhibitory polypeptide (GIP) have
the ability to induce glucagon secretion and can be stored in parasympathetic nerve
terminals. Galanin and neuropeptide Y is found in sympathetic terminals [168].
1.8 Diabetes
In the UK alone the number of people diagnosed with diabetes has reached 3.5
million and this figure is expected to rise to 5 million by 2025 making diabetes one of
the biggest health problems. The majority of cases (~90%) are patients with Type 2
41
diabetes (T2D). Globally, the prevalence of diabetes is expected to increase from 171
million in 2000 to 366 million by 2030 [175]. Diabetes is a consequence of the loss of
glycaemic control and islet dysfunction is a characteristic of both Type 1 (T1D) and 2,
as defined in Sections 1.8.1 and 1.8.2 below. It was 40 years ago that the bihormonal
hypothesis to explain the diabetes pathophysiology was first proposed [176]. This
hypothesis postulated that the disease was caused by an insulin deficit or resistance
accompanied by absolute or relative glucagon excess which results in an elevated
rate of glucose production by the liver surpassing glucose utilisation and thus leading
to hyperglycaemia. Insulin deficiency is the major feature of diabetes and results in
chronic hyperglycaemia with glucose levels above 7 mM which can lead to long-term
health issues such as kidney failure, neuropathy, blindness and cardiovascular
disease [177].
1.8.1 Type 1 diabetes
Type I is caused by the T cell mediated auto-immune destruction of β-cells. In T1D
patients, islets become infiltrated by CD8+ T cells which appear to be exclusively
specific towards islet autoantigens [178]. This usually occurs in people with a genetic
susceptibility and is believed to be initiated by environmental stressors such as a
viral infection [179]. The major susceptibility locus is located at the HLA class II genes
[180]. Patients normally have islet autoantibodies and the infiltration of islets by
immune cells results in an overall reduction of 70-90 % of β-cell mass at clinical onset
with most islets having lost their β-cell element [181].
1.8.2 Type 2 diabetes
This form of disease has reached epidemic proportions in various parts of the world,
a result of adverse environmental components, particularly the increased
accessibility to high caloric food accompanied by a sedentary lifestyle and the lack of
sufficient levels of physical activity [182]. The heritability of T2D is well established,
and variations of specific genetic loci interact with such environmental factors which
predispose to disease and have an impact on the development of the diabetic
phenotype [183]. Loss of glycaemic control in this type of diabetes is a combined
42
result of defective insulin secretion from β-cells and insulin resistance of insulin
target tissues such as muscle, liver and adipose tissue [184]. Development of obesity
is an important risk factor foretelling the eventual development of insulin resistance
[185] which, in an individual with a genetic predisposition to β-cell impairment,
causes a change in glucose tolerance. In the initial stages of T2D patients maintain
their glucose tolerance by an increase in functional β-cell mass and hypersecretion of
insulin as a compensatory mechanism for insulin resistance. However, as the disease
advances β-cells are no longer able to secrete enough insulin to reach the levels
required for peripheral tissues and that is when glucose intolerance begins [184].
Hyperglycaemia in the fasting and fed state, a consequence of dysregulated glucose
production by the liver and diminished glucose clearance, is attributed to ineffective
insulin action. A role for glucagon in the pathogenesis of T2D and a bihormonal axis
of the disease has long been suspected and multiple studies now support this view
[186].
1.8.3 Monogenic diabetes
The prevalence of monogenic diabetes is thought to be at 2-5% of the diabetic
patient population [187]. It represents a heterogeneous set of disorders caused by
faults in single genes. The monogenic variants are categorized based on the age of
onset, neonatal diabetes appearing before six months of age and maturity onset
diabetes of the young (MODY) showing signs before 25 years of age, with an
autosomal dominant type of inheritance and insulin independence [188]. MODY can
be caused by mutations, deletions or other abnormalities in any of at least twelve
different genes resulting in intrinsic β-cell dysfunction. These genes, which include
glucokinase [189], the transcription factors HNF-4α [190], HNF-1α [191], HNF-1β
[192], IPF1 [193] and NeuroD1 [194], are all expressed in β-cells.
1.9 Genetics of T2D
Although the global epidemic of T2D is largely driven by lifestyle and nutritional
changes, the blending of environmental factors with susceptible genetic traits
promotes the development of the disease [195]. For T2D patients diagnosed in
43
adulthood, the concomitant action of such susceptibility alleles and several
combinations of frequent variants at many loci could lead to detrimental effects
upon interaction with the aforementioned predisposing environmental components
[182].
The concept of a genetic influence on T2D is now commonly acknowledged. A study
on young patients with T2D revealed that the majority of the parents were either
diabetic or at least glucose intolerant. Similarly, there was high prevalence of
diabetes or glucose intolerance in the siblings, reaching about 70% [196]. The high
prevalence of diabetes in particular ethnic groups provides further evidence of a
genetic component [197]. Studies on monozygotic twins revealed that when one was
diagnosed with diabetes 48 out of the 53 pairs were concordant, with the majority of
the second twin in each pair becoming diabetic within five years of the first [198].
The ability to scan the whole genome for variation and do so using high throughput
technology made it possible to search for genetic variation to explain phenotypic
diversity and disease risk in genome-wide association studies (GWAS) (Section 1.9.2).
Assaying single nucleotide polymorphisms (SNP) to study the association of genetic
variation and disease has now become the norm and can give clues into the genetic
architecture contributing to complex disease [199].
1.9.1 Single nucleotide polymorphisms (SNPs)
SNPs are single base pair changes in the DNA sequence that are abundant in the
human genome [200]. In the case of genetic studies, SNPs are normally used as
markers of a part of the genome with the vast majority of them having no or minor
effect on the biology of an individual. However, SNPs can have a functional impact, a
consequence of causing amino acid changes, or, in the case of SNPS in non-coding
(intergenic or intronic regions), altering transcript stability or transcription factor
binding ability [201]. SNPs are the most frequent type of genetic variation in humans
[202] occurring on average every 300 base pairs (bp). Normally, there are two forms
of each SNP (one per haploid chromosome) so that in any given population, two
naturally occurring base pairs will likely exist for each specific SNP location with each
44
individual being either heterozygous or homozygous for an SNP. The major allele is
the most common variant of the SNP, while the minor the one with the lower
frequency.
1.9.2 Genome wide association studies (GWAS)
Although linkage analysis and candidate gene approaches have in the past been
successful in revealing familial genetic defects with strong effects and have allowed
the identification of genes that cause MODY, these approaches are not effective in
identifying common genetic variants which have a relatively moderate effect on
disease risk [199].
In GWAS studies, the genome of patients and non-patients is compared to find SNPs
that are associated with disease. Through the use of microarrays a staggering
number of 500 000 to a million SNPs are genotyped during each study, allowing for a
much faster discovery of risk variants linked to T2D. These kinds of studies can lead
to the discovery of several gene variants, each individual one having a small impact
on disease risk. When a particular SNP is found associated with disease, the locus is
typically annotated with the name of the nearest gene. Despite this, it should not be
automatically assumed that the variant involved is the molecular defect that
accounts for the disease phenotype or that the closest gene is necessarily implicated
with disease. The annotation merely marks a specific part of the genome that
contains the variant which could actually be functioning at a distance for example
regulating gene expression at a distant site [199].
The first GWAS for T2D was performed on a French case-control cohort, comprising
661 T2D cases and 614 healthy controls testing a total of 392,935 SNPs for
association with the disease. This study led to the identification of novel and
reproducible association signals at the SLC30A8 and HHEX loci [203] and at the same
time confirmed the already known association at TCF7L2 [203]. Shortly afterwards, a
study on an Icelandic population, confirmed the loci just mentioned and identified a
45
new association that of a CDKAL1 variant with an odds ratio (OR) of 1.20 [204].
Further replication was provided by Zeggini et al [205].
An OR is a statistical term in this case used to quantify how strongly a specific allele
is associated with disease. An OR significantly greater than 1 would mean an allele is
associated with disease. The strongest association was mapped to a variant in the
transcription factor TCF7L2 gene which had been previously identified in Icelandic
and Danish populations [206]. Each copy of the risk allele carries a high odds ratio for
T2D of 1.4-1.5 [207]. Inheritance of the risk allele can also be used to predict the
possibility of transitioning from prediabetic conditions to T2D [208].
Currently the model of T2D susceptibility in adulthood is that, being a complex
metabolic disease, it is probably the consequence of environmental issues on
predisposed individuals bearing a mixture of increased risk genetic variants. It is
believed that each risk allele, on its own, has a moderate effect with odds ratios of
1.10-15 [182].
Ever since the initial investigation [203] several others have undertaken such studies,
confirming previously identified disease signals or discovering new T2D associated
variants [205, 209]. Although a lot of the common variant signals revealed through
GWAS are associated with impaired islet function, the majority of the signal from the
studies is in non-coding regions and some in introns [210]. Determining functional
links to specific genes is thus more challenging requiring the deployment of both
mapping approaches [211] and the use of “functional genomics” to test directly the
impact of forced changes in the expression of candidate genes in model systems
(cells, animals etc) [212].
1.9.3 SLC30A8 and T2D risk
One of the first variants to be discovered by GWAS, in the initial study by Sladek et al
[203], was rs13266634, a non-synonymous SNP in SLC30A8 located in a 33 kb linkage
disequilibrium block on chromosome 8, which includes only the 3’ end of the gene
[203]. The SNP causes an amino acid change from tryptophan to arginine at position
46
325 at the C-terminus of the ZnT8 protein. Exon 13, encoding the last 52 amino
acids of both ZnT8 isoforms contains a total of 6 SNPs, with a second SNP occurring
at position 325, rs16889462, in this case changing the amino acid from an arginine to
glutamine. The gene product ZnT8 is expressed primarily in pancreatic islets [213]
where it is likely to play an important role in islet hormone secretion.
The rs13266634 allele was associated with increased T2D risk. Possession of only one
allele of the risk variant results in an OR of 1.18, while being homozygous, increases
the OR to 1.53 [203]. Subsequent studies performed on T2D patients from different
countries found similar association between SLC30A8 and disease [204, 214]. The
association of this SNP with T2D typically involves comparatively young onset and
leanness, although it was not identified as a cause of MODY following extensive
studies on 53 extended families of patients with early onset T2D [215]. Of note, ZnT8
was identified as a target by autoantibodies in 60-80% of new onset T1D patient
[216] though the risk variants discovered by GWAS do not affect T1D risk.
Studies performed on a German population of T2D patients [217, 218] revealed that
the risk SLC30A8 allele, which is also the major allele in most populations, is
associated with decreased insulin release following intravenous administration of
glucose and defective conversion of proinsulin to insulin during an oral glucose test,
whilst there were no signs of insulin resistance. Another study on a French
population [219] demonstrated that those with the risk allele had higher plasma
glucose levels in fasting conditions and decreased insulin secretion. However, other
studies showed no connection between the SNP and markers of β-cell function [214,
220].
Unexpectedly, a more recent study [221] showed that very rare loss of function
variants of SLC30A8 are actually protective and were associated with decreased T2D
risk. 12 protein truncating variants were identified and carriers of such variants had a
65% reduction in T2D [221]. Such observations conflict with the findings that
expression of a less active (R325) form of ZnT8 [222] results in an elevated risk of
47
T2D suggesting that a more intricate mechanism is at play regarding the rs13266634
T2D association [223].
1.10 The α-cell and glucagon secretion in T2D
Similar to the β-cell, the α-cell becomes dysfunctional in diabetes. Plasma glucagon is
significantly elevated in cases of uncontrolled T1D and contributes greatly to
ketoacidosis [224]. Glucagon levels are abnormal in T2D as well, although the
changes are more subtle, tending towards enhanced secretion and/or insufficient
inhibition of α-cells in most patients. Such inappropriately high glucagon levels
appear to have a role in hyperglycaemia and impaired glucose regulation [186]. In
conditions of inadequately-controlled diabetes or ketoacidosis, plasma glucagon
levels are very high [224]. Hyperglucagonaemia in the fasted state has been
observed in some but not the entirety of T2D patients with at least moderate
glycaemic control [225].
In the diabetic islet, the α-cell does not respond appropriately to raised plasma
glucose. The suppression of glucagon secretion as seen in healthy individuals by
elevation of glucose levels does not occur in T2D. The response to hyperglycaemia is
either blunted or lacking and plasma glucagon stays abnormally high at similar blood
glucose concentrations [186]. Additionally, stimulation of glucagon release by
normal stimuli including intravenous arginine or protein-rich meals is greater in T2D
compared to healthy individuals [226]. The inability of hypoglycaemia to induce
glucagon secretion is well established in T1D, but whether this is the case in T2D is
more controversial. Several groups have found the counter-regulatory response to
hypoglycaemia by the α-cell to be weakened at least in individuals with long duration
of T2D [227, 228].
Fasting hyperglycaemia in T2D is directly attributed to abnormal hepatic glucose
production and hepatic resistance to suppression of glucose production by insulin is
evident in patients [186]. However, surplus glucagon in the fasting state could also
be a driver for inappropriate levels of glucose production by the liver. Basal glucagon
release is a significant factor in the control of fasting glucose levels in both diabetic
48
and healthy individuals [225, 229]. Fasting glucagon can be up to 50% higher in some
T2D patients compared to non-diabetics [225].
In order to determine the importance of glucagon in diet-induced and genetic T2D in
a rodent model, as caused by leptin receptor (LepR) ablation, one group studied high
fat diet-induced obese (DIO) mice and LepR KO (LepR-/-) mice with or without
glucagon receptors (GcgR) [230]. GcgR+/+ and GcGR-/- mice were placed on a high
fat diet and, as previously observed, the body weight of the KO mice did not increase
substantially and they did not exhibit hyperinsulinaemia or hyperglycaemia in
contrast to the WT mice. DIO and LepR-/- mice with functional GcgRs were both
hyperinsulinaemic and obese developing mild and severe T2D respectively. LepR-/-
GcgR-/- mice showed no signs of hyperinsulinaemia and hyperglucagonaemia.
Delivering GcgR to the liver through the use of an adenovirus led to the severe
hyperinsulinaemia and hyperglycaemia normally present in LepR-/- mice, while
spontaneous disappearance of the transgene reversed the T2D phenotype. Such
findings indicate that glucagon is essential for hyperglycaemia in T2D and that it
enables diet-induced hyperinsulinaemia. These effects can be prevented by blocking
glucagon action.
1.11 Zn2+ in diabetes
A total of 2-3 g of Zn2+ can be found throughout the human body and this ion is most
abundant in the pancreas which contains 140 μg/g [231] with the highest
concentration within islets. In the plasma total Zn2+ reaches 10-20 μM and its free
concentration is in the nM range [232]. A role for Zn2+ in diabetes pathophysiology
was discovered almost 80 years ago, when a 50 % reduction in pancreatic Zn2+ was
observed in the diabetic compared to non-diabetic cadavers [233]. Then, about 30
years later, studies in rodents revealed that a Zn2+ deficiency through diet led to a
decrease in glucose-induced insulin secretion and insulin levels in the islet [234].
Following that, it was shown that Zn2+ deficiency also causes a reduction in β-cell
granulation [235]. It has been reported that T2D patients exhibit a substantial
reduction in total blood Zn2+ as well as at the cellular level in comparison to healthy
49
controls accompanied by excessive zincuria [236]. Dietary supplementation
improved glucose handling in human subjects [237].
Research on various genetic mouse models of T2D demonstrated decreased
pancreatic Zn2+ levels [238]. Supplementation of db/db mice with Zn2+ led to
normalization of pancreatic Zn2+ concentration and a lessening of hyperglycaemia
and hyperinsulinaemia [239]. The same experiment in ob/ob mice increased
pancreatic insulin levels and similarly reduced the observed fasting hyperglycaemia
and hyperinsulinaemia as well as the excessively high insulin release in response to
glucose in vitro [238]. The opposite experiment of a Zn2+-deficient diet in db/db mice
caused an exaggeration of fasting hyperglycaemia [239].
1.12 Cellular Zn2+ homeostasis
Zn2+ is very important biologically having catalytic, structural and regulatory roles
[240]. Additionally, it is involved in a plethora of cellular processes and is believed to
be a cofactor for approximately 3000 proteins in the human genome including
enzymes, transcription factors and hormones [241]. Mechanisms that regulate Zn2+
uptake, distribution and efflux are crucial for maintaining homeostasis and thus
cellular function. Intracellular Zn2+ homeostasis is maintained in part through the
activity of Zn2+ binding proteins and a variety of Zn2+ transporters belonging to two
protein families which have opposing function but work in a coordinated way (Figure
1.5).
1.12.1 Metallothioneins
The metallothioneins (MTs) are intracellular Zn2+-binding proteins essential in
regulating Zn2+ uptake distribution, storage and release. They are redox active,
having sulphur donors, controlling Zn2+ homeostasis but also contributing to the
redox state of the cells by releasing Zn2+ in the presence of oxidative stress [242].
The capacity of MTs for intra- and intermolecular Zn2+ exchange means that MTs
keep sufficient Zn2+ accessible in the cell at micromolar concentrations, higher than
the concentration of free Zn2+ in the picomolar range, which would be too low to act
as a reservoir of Zn2+ for Zn2+ proteins [243]. Several SNPs in the MT genes have been
50
discovered to be associated to T2D following studies done on an Asian population
[244].
1.12.2 ZIP transporters
The ZIP (SLC39A) family of Zn2+ transporters has 14 members (Zip1-14) in mammals.
The majority are predicted to have eight transmembrane domains and a long loop
with a histidine-rich region which is likely to be the Zn2+ -binding domain. They have
a similar predicted topology with both their N and C termini located on the
extracellular surface of the plasma membrane [245]. Their function is the facilitation
of Zn2+ influx from either the extracellular space or within intracellular
compartments into the cytoplasm, thus increasing the cytoplasmic free Zn2+
concentration.
1.12.3 ZnT transporters
The ZnT (SLC30A8) family comprises 10 members, ZnT1-10, and most are predicted
to be made up of six transmembrane domains with cytoplasmic carboxy and amino
termini. Similarly to the Zips, they contain a long histidine-rich loop potentially being
the site of Zn2+ binding in the cytosol [246]. ZnT5 differs in that it has 15
transmembrane domains [247], whilst ZnT6 has a serine rich loop instead [248]. The
majority of ZnTs form homodimers, except ZnT5 and ZnT6 which form heterodimer
responsible for transporting Zn2+ early in the secretory pathway [249]. ZnTs have
been found in cytoplasmic compartments normally associated with the Golgi
complex, endosomes or endoplasmic reticulum [250]. In contrast to Zips they are
efflux transporters, carrying Zn2+ from the cytosol into intracellular compartments or
extracellularly.
Zn2+ binds to the histidine rich region of the protein and the transmembrane
domains form the pore for Zn2+ transport in both transporter families [246]. The
different isoforms vary in their affinity for Zn2+ and also have the ability to transport
other transition metals, for example manganese, cadmium, iron and copper which
51
may inhibit the transport of Zn2+ [251]. Transcription and translation of at least some
of these transporters appears to be regulated directly by Zn2+ [251].
Some members of the ZIP and ZnT families have ubiquitous expression but others
are tissue specific. For example ZnT1, ZnT5 and Zips 6-10 are found throughout the
body, but ZnT10 is only expressed in the liver and brain [251].
1.13 ZnT8
ZnT8 was first identified as a pancreas-specific ZnT member and cloned by Chimienti
et al in 2004 [213]. It was predicted to adopt the same topology at the other ZnT
proteins, having 6 transmembrane helical domains with a histidine rich region
between helices 4 and 5. These authors showed that only the pancreas had
significant expression of ZnT8 and that although not complete there was a high
degree of colocalization between insulin and EGFP-tagged ZnT8. The protein was
detected in intracellular vesicles and at the plasma membrane. Additionally, using
Zinquin a Zn2+ indicator that localizes to intracellular vesicles [252], ZnT8 was found
to be involved in Zn2+ transfer across the vesicle membrane following transient
expression in HeLa cells which do not normally express the transcript. Since co-
staining for ZnT8 with insulin granules was not complete, ZnT8 could possibly be
localised at other subcellular sites and so it was found to partially colocalize with the
endoplasmic reticulum marker calnexin.
The human SLC30A8 gene is found on chromosome 8, contains eight exons across
37kb and is translated into a 369 amino acid protein. Mouse and rat ZnT8 have been
shown, respectively, to share 80 and 76% identity with the human form of ZnT8 and
are most closely related to ZnT2, ZnT3 and ZnT4 of the ZnT family. The greatest
homology is with ZnT2, reaching 53.5% [253]. Initially ZnT8 expression in the mouse
was found to be largely restricted to the pancreas, specifically in the α- and β-cells of
the islet [130]. Similarly analysis of a large number of human tissues confirmed the
pancreas specific expression of ZnT8. Later reports showed its expression, albeit at
low levels, in human adipose tissue [254], lymphocytes [255] and the cubical
epithelium lining thyroid follicles and the adrenal cortex [256].
52
Figure 1.5 – Subcellular localization and direction of transport of the ZnT and ZIP Zn2+
transporter families. The direction of Zn2+ transport is shown by arrow for the ZnT (green)
and ZIP proteins. From Myers et al [251].
53
Upon SDS-PAGE gel analysis ZnT8 migrates as bands of ~ 40 and 90 kDa, indicating
that it may form dimers. X-ray crystallography on the E.Coli Zn2+ transporter, a
bacterial ZnT8 homologue Yiip, showed that it is a homodimer bound in a parallel
orientation by co-ordinating to four Zn2+ [257]. The amino acid at position 325 in the
ZnT8 sequence is predicted to reside at the dimer interface. Nicolson et al [222]
demonstrated that the risk variant has a reduced ability to transport Zn2+ across the
granule membrane, with the position of the changed amino acid believed to be in a
different position to the one mentioned above, in a region near the predicted site of
Zn2+ binding or docking sites to Zn2+-binding proteins. Introducing positive charge
into this area through the presence of arginine instead of tryptophan might possibly
affect the kinetics of Zn2+ transport and the recruitment of Zn2+ binding proteins
[258]. Similar findings, supporting this view, were later made by Kim et al [259] using
radiotracers to monitor Zn2+ fluxes into isolated secretory granules. On the other
hand Weijers [260] has presented an alternative structure prediction suggesting that
the R-W325 replacement has little effect on structure or interactions with Zn2+ or
binding proteins. A resolution of this issue will require crystallisation of the ZnT8
protein itself (or the C-terminal domain) as well as further functional assessment of
the impact of the common polymorphism on activity.
There is not a lot of information on the transcription factors involved in the
regulation of ZnT8 expression. A strong transcriptional enhancer has been
discovered in the second intron of the mouse gene [261] which is highly conserved in
humans [262]. It was found that this binds several transcription factors one of them
being Pdx1. However this enhancer is only active in β-cells and not α-cells. In terms
of factors that could modulate its expression, Zn2+ deprivation, glucose and cytokines
have been shown to reduce Slc30A8 expression in both mouse and rat cell lines [263,
264] though no such effect was observed on human islets after administration of
cytokines [265].
54
1.13.1 ZnT8 and the β-cell; studies using mouse knockout lines
Zn2+ is essential in the β-cell for the correct processing and storage of insulin in
secretory granules [266]. After translation proinsulin is transferred to the Golgi body
where Zn2+ is abundant and there proinsulin hexamerizes with Zn2+, two ions in each
hexamer. In the immature secretory vesicle, proinsulin in processed into insulin and
insulin crystallization happens in the insulin granule, creating the characteristic
dense core [267].
Thus given this importance of Zn2+ in insulin granules and since ZnT8 was discovered
to be a T2D gene, there was a lot of interest into its role in the β-cell. Functioning
across the granule membrane to transport Zn2+ into the lumen for insulin
crystallization, ZnT8 has the potential of being extremely important for the β-cell.
The role of ZnT8 in the β-cell has been examined with either global or β-cell specific
deleted mouse strains. The results from these studies were controversial, with not
everyone agreeing with the findings of the other, although there were some
concurring points. None of the groups observed any changes in total insulin content,
islet size or in the proportion of cell types [222, 268-271]. In the majority of the
cases, whether global or β-cell specific ablation, the insulin granules appeared to
have been changed morphologically [222, 270-272]. Only one study reported no
changes [269]. Reductions in cellular Zn2+ content were also reported though there
were some differences between groups. One group observed a reduction in Zn2+
content with the use of Zinquin and dithizone yet not with FluoZin-3 [222], perhaps
reflecting the localisation of these dyes to the granules (zinquin and dithizone) and
cytosol (FluoZin-3). On the other hand, Pound et al did demonstrate a significant
decrease in cytosolic free Zn2+ using FluoZin-3 [268, 269]. Additionally another group
only showed a modest reduction in cytosolic Zn2+ when looking at β-cell specific KO
mice versus controls [272] compared to the changes observed in global KO mice
[222]. Of note the fluorescent intracellular probes used above provide uncertain sub-
cellular localisation and may be accumulated into organelles including secretory
granules to different degrees.
55
Glucose-stimulated insulin secretion from isolated islets was one of the most divisive
findings since two groups described an impairment of secretion [268, 272], two
elevated secretion [222, 271] and two no effect at all [269, 270]. It has been
suggested that the increase in insulin release might be due to the alleviation of
inhibition by Zn2+ which was previously reported [271].
Zn2+ is important for the maturation of the enzymes involved in proinsulin processing
and so proinsulin levels and processing were investigated. Again the results were in
disagreement, with two studies detecting no changes in proinsulin processing [222,
270], two showing a small increase in plasma levels [271, 272] while Pound et al
[269] observed a reduction in plasma proinsulin levels which was, however, affected
by genetic background. Thus it appears that despite the marked changes in granule
morphology the deletion of ZnT8 has minor effects on proinsulin processing per se.
All of the groups above examined the effect of ZnT8 deletion on in vivo glycaemic
parameters. No effect of ZnT8 deletion on insulin sensitivity was reported and no
change in fasting blood glucose except by one group who showed an increase but
only in young mice [222]. Three studies observed no changes in fasting insulin [269]
[271, 273] and only one detected a decrease [222]. However, most studies showed
impaired glucose tolerance during intraperitoneal glucose tolerance tests in young
mice [222, 269-271] with only two reporting such an effect in old mice [222, 268-
271]. Lastly, one group reported impaired glucose tolerance assessed by oral glucose
tolerance test in young mice [269], but not in older ones [272].
Studies in the INS-1 rat β-cell line using an shRNA-mediated approach to eliminate
ZnT8 expression by >90% resulted in reduced uptake of exogenous Zn2+, lowered
insulin content and decreased insulin secretion supporting the view suggesting that
ZnT8 may have a role in insulin release and the control of glycaemia [263].
56
1.13.2 ZnT8 and the α-cell so far
The presence of ZnT8 in the α-cell has been confirmed by qPCR and by
immunocytochemistry [222]. Just as in the β-cell, colocalization of ZnT8 with
glucagon was only partial with ZnT8 found in other cytosolic locations other than the
secretory granules. The function of ZnT8 in the α-cell has not been extensively
studied. Only one group has previously provided a limited amount of data on a
αZnT8 KO mouse [222]. Intraperitoneal glucose tolerance tests showed that the
absence of ZnT8 had no impact on glucose homeostasis. Similarly, there were no
differences in plasma glucagon levels when compared to control mice [272].
However, these studies were confined to mice of a specific age, and no other
parameters were tested.
There have been some studies done on the effect of overexpression or knockdown
of ZnT8 on αTC1.9 cells. Overexpression of either the R325 or W325 variants in these
cells caused a reduction in glucagon content and a decrease in glucagon secretion by
50%. Knockdown of ZnT8 reciprocally led to an elevation of glucagon mRNA and
increased glucagon secretion by 70% [274].
1.14 Targeting glucagon for T2D treatment
Taking into consideration the scientific evidence supporting glucagon as a
contributing factor in abnormal glucose homeostasis in T2D, targeting glucagon
action could be a potential therapeutic strategy.
Reducing plasma glucagon levels using somatostatin, while insulin levels are kept
constant, was found to normalise the enhanced basal rate of hepatic glucose
production and fasting plasma glucose concentration in hyperglycaemic patients
with T2D [225]. Glucagon receptor antagonists have been developed which, in
leptin-deficient diabetic mice, have been demonstrated to be effective in lowering
the enhanced basal rate of hepatic glucose production and fasting plasma glucose
levels [275].
57
Orally administered glucagon receptor antagonists have also been produced but the
concern regarding this type of drugs is the ensuing α-cell hyperplasia and the
constant elevation of blood glucagon concentration [276]. There is also evidence,
however, that glucagon receptors are required in the liver to prevent hepatic lipid
accumulation [277], a significant counter-indication in T2D. However, drugs which
cause a reduction in plasma glucose concentration without severe α-cell
hypertrophy and only a modest elevation in blood glucagon levels have been
developed [278]. Two of these glucagon receptor antagonists had reached phase 2
trials and have exhibited robust haemoglobin A1c-reducing efficiency [279], yet the
studies were discontinued due to an elevation in LDL cholesterol, presumably
reflecting the requirement for hepatic glucagon signalling mentioned above.
Aims of the thesis
The role of SLC30A8 in the α-cell has not been characterized. The aim here was to
investigate the effect ZnT8 has on glucose homeostasis and glucagon secretion. In
order to do this mice were generated with α-cell specific deletion or overexpression
of ZnT8.
58
Chapter 2
Materials & methods
59
2.1 In vivo animal work
2.1.1 Mice maintenance and generation
Animals were kept in a pathogen-free facility under a 12 h light-dark cycle with
access to water and a standard mouse diet (Lillico Biotechnology). After weaning at
3-4 weeks of age they were housed two to five per separately ventilated cage. The
transgenic mouse strains were maintained on a C57BL/6 genetic background. All
procedures were performed in accordance with UK Home Office regulations under
the Animals (Scientific Procedures) Act 1986 (PPL 70/7349, Dr Isabelle Leclerc).
2.1.2 Generation of transgenic mice
Mice bearing SLC30A8 alleles where exon 1 was flanked by LoxP sites were obtained
from GenOway (Lyon, France). To generate mice with ZnT8 deleted specifically in the
α-cell ZnT8fl/fl mice [272] were crossed to mice bearing the Cre transgene under the
control of a ~0.6 kb fragment of the pre-proglucagon promoter (PPGCre) [280].
2.1.3 Glucose tolerance tests
Mice were fasted overnight before intraperitoneal injection with glucose (1g/kg).
Blood samples were subsequently taken from the tail vein at 0, 15, 30, 60 and 120
min and glucose concentration was measured with an automatic glucometer as
previously described [222].
2.1.5 Insulin tolerance tests
Mice were fasted for 5 h before intraperitoneal injection with insulin (0.5U/kg).
Blood samples were subsequently taken from the tail vein at 0, 15, 30 and 60 min
and glucose concentration was measured with an automatic glucometer as above.
Additionally, plasma (~100 μl) was taken at time 0 and 15 min for glucagon
measurements using a radioimmunoassay (Millipore).
60
2.1.6 Fasting plasma glucagon measurements
Mice were fasted overnight before taking blood samples of about 40 µl from the tail
vein. Plasma was separated from the cellular components by centrifugation at 12000
rpm for 10 min at 4 °C. A glucagon RIA kit (Millipore) was used to measure glucagon.
2.1.7 Streptozotocin treatments
Mice at the age of 6-7 weeks were treated with 50 mg/kg streptozotocin in a 0.1M
sodium citrate buffer pH 4.5 for five consecutive days following an overnight fast.
Fasting hyperglycaemia (14-19 mM) was evident about 4 weeks later and IPGTT were
performed at this point.
2.1.8 Hypoglycaemic clamps
The clamps were performed by S. Migrenne, C. Magnan (University Paris Diderot-
Paris 7-Unit of Functional and Adaptive Biology (BFA) EAC 7059 CNRS, France) as
described [281].
2.2 In vitro experiments
2.2.1 Islet preparation
Animals were terminated in a CO2 chamber, and death was confirmed by cervical
dislocation. The abdomen was opened along the sternum for extraction of the
pancreas. Isolated pancreata, inflated with collagenase solution at 1 mg/ml (Serva),
were placed in a water bath at 37 °C for 10 min. Cold (~4°C) additive-free RPMI
(Sigma) was added followed by centrifugation at 1000 rpm for 1 min. The
supernatant was removed before adding about 15 ml of cold additive-free RPMI and
the pancreas was resuspended in the media. This pellet was washed three more
times. The islets were then resuspended in 3 ml of Histopaque 119 (Sigma). A
sucrose gradient was then made by adding dropwise 3 ml of Histopaque 1083 and
1077 (Sigma). Finally, RPMI was added dropwise up to 12 ml. The gradient was
61
centrifuged at 2500 rpm for 20 min. The islets found just below the RPMI layer were
collected and centrifuged at 1000 rpm for 1 min after addition of up to 12 ml RPMI.
The supernatant was removed; islets were resuspended and placed in a Petri dish
containing 15 ml complete RPMI media. After a few hours of recovery at 37 °C islets
were handpicked and placed in 15 ml of fresh media.
2.2.2 Islet dissociation
After overnight culture islets were hand-picked and washed once in 5 ml phosphate
buffered saline (PBS). Then 3-4 ml of Hank’s dissociation buffer (Invitrogen)
supplemented with 0.01% BSA, was added and the islets were centrifuged at 2000
rpm for 2 min. This was repeated once more. The islets were then dissociated using a
P200 pipette in 150 µl of Hank’s buffer and 20 µl 0.1% trypsin solution (Sigma).
Following dissociation, 20 µl of fetal bovine serum (FBS) was added and the cells
were washed with PBS and either resuspended for plating on coverslips coated with
polylysine in 30-40 µl media per coverslip or were incubated with DAPI for 10 min to
stain for dead cells prior to FACS.
2.2.3 Glucagon secretion
Experiments were performed in 12 well plates. Size-matched islets (18) were placed
in 1 ml of 10 mM glucose solution for 30 min in a 37 °C water bath with continuous
and gentle agitation at ~70 revolutions per min. The buffer was then replaced with
fresh KRBH containing either 1 mM or 17 mM glucose for further incubation at 37 °C
for 1 h. The supernatant was collected (secreted glucagon) and the islets were lysed
by sonication for about 10 s in acidified ethanol (500 μl; total glucagon). Secreted
and total glucagon were measured by radioimmunoassay (Millipore). Samples (100
μl) were used for the assay. Sample was mixed with anti-glucagon antibody at 4 °C
for 20-22 h before addition of I125 glucagon for 22-24 h again at 4 °C. Precipitating
agent (Goat anti-Guinea Pig IgG Serum, 3% PEG and 0.05% Triton X-100 in 0.05M
Phosphosaline, 0.025M EDTA, 0.08% Sodium Azide) was then added for 20 min at 4
62
°C and centrifuged at 4000 rpm for 20 min. The standard curve, total and non-
specific binding values were used to calculate the glucagon content of secreted and
total samples. Secreted glucagon was expressed as a percentage of total glucagon.
2.2.4 Ca2+ imaging with Fluo-2
Islets were incubated (37 °C 95% O2/5% CO2) for 45 min in 10 μM Fluo-2-AM
(Invitrogen) dissolved in DMSO (0.01% wt/vol) and pluronic acid (0.001% wt/vol) in a
bicarbonate-based buffer (120 mM NaCl, 4.8 mM KCl, 1.25 mM NaH2PO4, 24 mM
NaHCO3, 2.5 mM CaCl2, 1.2 mM MgCl2) pH 7.4 containing 3 mM glucose. Following
incubation, 3-4 islets were placed in a perfusion chamber, mounted on a Zeiss
Axiovert confocal microscope and perfused continuously at 37 °C with buffer
containing 1mM glucose followed by 11 mM glucose. Fluo-2 was excited with a 491
nm laser and emitted light was filtered at 525/50 nm. VolocityTM was used for data
capture and ImageJ [282] for image analysis. Traces are shown as normalized
intensity over time (F/Fmin).
2.2.5 Zn2+ imaging with FluoZin-3
Islets were incubated (37 °C 95% O2/5% CO2) for 1 h in 1 μM FluoZin-3-AM
(Invitrogen) dissolved in dimethyl sulfoxide (DMSO) (0.01% wt/vol) in a bicarbonate
buffer containing 11 mM glucose. Islets were transferred into a perfusion chamber
at 37 °C and continuously perifused with buffer. Imaging was done at baseline and
after addition of the Zn2+ chelator N,N,N′,N′-tetrakis(2-pyridylmethyl)
ethylenediamine (TPEN) using the Zeiss Axiovert confocal microscope. FluoZin-3 was
excited with a 491 nm laser and emission monitored at 525/550 nm. Results were
presented as F1/Fmin where F1 was mean baseline fluorescence and Fmin minimum
fluorescence in the presence of the Zn2+ chelator. Traces were analysed using an
Image J macro.
63
2.2.6 Granular Zn2+ imaging with Zinpyr-4
Islets were incubated as above for 30 min in 10 μM Zinpyr-4 (Santa Cruz) [283].
Imaging was performed on a Nikon Ti Eclipse microscope equipped with an X-Light
spinning disk and Hamamatsu ORCA-Flash4.0 CMOS detector. Static images were
captured at 37 °C using a 63x objective, keeping the exposure time and laser
intensity constant throughout. Zinpyr-4 and RFP were excited using 488 nm and 561
nm lasers, respectively, and signals monitored at 535/50 nm (Zinpyr-4) and 630/75
nm (RFP). Differences in Zinpyr-4 intensity between WT and KO animals was
measured using the corrected total cell fluorescence (CTCF) (integrated density –
(cell area x mean background fluorescence)), as described [284].
2.2.7 Mammalian cell culture
αTC1.9 cells, a mouse glucagonoma cell line provided by the American Tissue Culture
Collection (ATCC) [285], were grown in Dulbecco’s modified Eagle’s medium (DMEM)
without glucose (Sigma) supplemented with 10% FBS, 15 mM Hepes, 0.1 mM non-
essential amino acids, 0.02% BSA, 1.5 g/L sodium bicarbonate, 2 g/L glucose and 1%
penicillin/streptomycin in a 37 °C incubator with 95% O2/5% CO2.
INS-1 (832/13) cells [286], a rat insulinoma cell line, were grown in RPMI medium
(Sigma) containing 11mM glucose, supplemented with 10% FBS, 2 mM glutamine,
1% penicillin/streptomycin, 10 mM Hepes, 1 mM sodium pyruvate and 50 µM β-
mercaptoethanol in a 37 °C incubator with 95% O2/5% CO2.
HEK 293 cells [287], a human embryonic kidney line, were grown in DMEM
containing 25 mM glucose, supplemented with 10% FBS, 1% penicillin/streptomycin
in a 37 °C incubator with 95% O2/5% CO2.
64
2.2.8 Zn2+ imaging with eCALWY-4: cells
αTC1.9 cells were plated on 24 mm coverslips and cultured overnight to grow. The
following day they were infected with the eCALWY-4 adenovirus, a Förster
resonance energy transfer (FRET) based sensor [288], at a multiplicity of infection of
25-30 for imaging the next day. Cells on coverslips were washed twice in Krebs
Hepes-bicarbonate (140 mM NaCl, 3.6 mM KCl, 0.5 mM NaH2PO4, 0.2 mM MgSO4,
1.5 mM CaCl2, 10 mM Hepes, 2 mM NaHCO3) buffer pre-conditioned with 95:5
O2:CO2 at a pH of 7.4. Solutions of TPEN in ethanol and pyrithione in DMSO were
prepared on the day of the experiment and used at a final concentration of 25 mM
and 1 mM respectively. Cells were maintained at 37 °C using a heating stage and
perifused with Krebs Hepes buffer initially with no additions. Calibration of the FRET
probe was performed by perfusing with the Zn2+ chelator TPEN followed by a
combination of Zn2+ (100 µM) and the Zn2+ ionophore pyrithione. The acquisition of
images was done with a 100 ms exposure time and 433 nm excitation. The
microscope was fitted with a monochromator and for FRET measurements a dichroic
mirror and two emission filters had to be used. Cells were imaged with a 40x oil
immersion objective.
The FRET-based probe undergoes a large ratiometric change following Zn2+ binding
[288]. The probe has a donor and an acceptor molecule and a conformational change
in the sensor domain results in a change in the ratiometric fluorescence signal from
these molecules. Dissociation of the two molecules means the donor emission is
detected upon excitation while when the two are in proximity the acceptor emission
is the one primarily detected because of FRET from the donor to the acceptor. In
eCALWY-4 the metal binding domains are Atox-1 and WD-4 linked to the
fluorophores Cerulean and Citrine respectively [288]. The metal binding domains are
linked by an amino acid linker. At low Zn2+ levels the metal binding domains are
separated from each other so that the two fluorophores can be close to each other
for generation of energy transfer. When Zn2+ levels increase, the metal binding
65
domains are in closer proximity separating the two fluorophores thus reducing
energy transfer (Figure 2.1).
Figure 2.1 – eCALWY sensor design. Atox1 and WD4 were used as metal binding domains.
Changes in conformation upon Zn2+ binding cause in energy transfer and thus fluorescence
that can be used to determine Zn2+ levels. From Vinkenborg et al [288].
2.4.4 Amplification and purification of adenoviral eCALWY4
AD293 cells (ATCC) were infected with 1 μl of the virus. After three rounds of
amplification the cells were harvested and were subjected to a freeze-thaw cycle to
release the virus. The cells were centrifuged at 5000 rpm at 4°C and the supernatant
was collected. Saturated CsCl (5.4 ml) in PBS was added to 9 ml of supernatant in a
13.5 Sorvall ultracentrifuge tube and was centrifuged for 10-16h at 4 °C at 65 000
rpm. The viral band was removed with a 23G needle, in dialysis tubing and placed in
500 ml dialysis buffer at 4 °C. The buffer was changed every hour 4 times. The virus
was then stored in storage buffer in 1:2 dilution in 40 μl aliquots at -80 °C.
2.4.5 Virus titration
AD 293 cells were plated ~24 h prior to infection on a 24-well plate. The cells were
infected with the eCALWY-4 adenovirus in serial dilutions when they reached ~70 %
confluency. After 24 h infection the cells were imaged with a 20x objective using an
66
Olympus epifluorescence microscope, counting the total number of green cells per
well which was used to calculate the MOI.
2.2.9 Immunostaining: cells
Cells were plated onto coverslips and the following day culture media was removed
before washing three times with PBS. Cells were fixed in 4% paraformaldehyde (PFA)
(w/v) in PBS for 15 min at room temperature and then washed with PBS. Cells were
permeabilized with 0.1% Triton X-100 in PBS for 10 min and subsequently washed
with PBS. 1% BSA in PBS was used as a blocking solution for 30-60 min at room
temperature. Incubations with primary antibodies at the appropriate dilutions were
performed at 4 °C overnight or for 48 h using 1% BSA in PBS. After more washes in
PBS cells were incubated with secondary antibodies at a 1:1000 dilution for 1 h at
room temperature on a shaker (30 revolutions per min) Coverslips were mounted on
microscope slides (Thermo Scientific) using DAPI containing mounting medium
(Vector Labs).
2.2.10 Immunostaining: islets
Fixation of islets was done in 4% paraformaldehyde (PFA) for two hours at room
temperature or overnight at 4 °C. After washing with PBS islets were incubated with
primary antibodies in 5% BSA plus 0.1% Triton X-100 in PBS overnight or for 48 h at
4°C. After more washes in PBS, islets were incubated with secondary antibodies at a
1:1000 dilution for 1h at room temperature on a shaker. Islets were transferred onto
slides before adding a drop of mounting medium (Vectashied with DAPI) and a
coverslip on top. The islets were imaged with a 20x magnification objective using a
confocal microscope.
67
2.2.11 Immunostaining: slices
Freshly isolated pancreata were fixed in 10% formalin (w/v) overnight at 4 °C,
embedded in wax and sliced into 5 µm sections. They were then dewaxed for 20 min
in Histoclear (Sigma) and rehydrated by washing for 5 min in 100, 95 and 70%
ethanol and distilled water. The sections were treated with vectors antigen solution
(Vectors Labs) 3-4 times washing with distilled water in between treatments. They
were then washed twice in PBS and blocked in PBS containing 0.1% Triton X-100, 2%
BSA and 2% goat and donkey serum for 60 min at room temperature. Primary
antibodies (anti-guinea pig insulin 1:200 (DAKO), anti-rabbit glucagon 1:100 (Sigma))
in PBS were then applied on the slides for overnight incubation at 4°C in a moist box.
The next day, slides were washed 3 times in PBS before incubation with secondary
antibodies (488 goat anti-guinea pig 1:1000, 568 donkey anti-rabbit 1:500) in PBS at
room temperature. There were some final washes in PBS before adding a drop of
Vectashield with DAPI on the slide and covering with a rectangular coverslip. The
slides were flat at 4°C and allowed to dry. Imaging was done on a Zeiss AxioObserver
widefield using a 40x objective. The α:β cell ratio was determined by subjecting
every islet image to manual thresholding (Image J macro written by Dr Pauline
Chabosseau) and dividing the α- cell area by the β-cell area.
2.2.12 Quantification of pancreatic glucagon content
Pancreata were extracted from mice fasted for 5 h, split into two, weighed and
homogenized using a Polytron homogenizer in 5 mL acidic ethanol (1.5% HCl, 70%
ethanol). They were incubated overnight at 4°C and homogenized further by
sonication. Another overnight incubation followed and the samples were centrifuged
at 2000 rpm at 4 °C for 15 min. The solution was removed and neutralized with an
equal volume of 1 M Tris-HCl. The samples were then diluted and glucagon
concentration was measured using a Homogeneous Time Resolved Fluorescence
(HTRF) glucagon assay (Cisbio) in a PHERAstar reader. The total amount of glucagon
was then calculated as ng/mg of pancreas.
68
2.2.13 Electron microscopy
Islets (~100-150) were handpicked, washed in PBS and centrifuged at 500 rpm for 30
s in a 1.5 ml eppendorf. PBS was removed and the islets were fixed in 500 μl freshly
prepared Vincenzo’s fixative (2% PFA, 2.5% glutaraldehyde, 3 mM CaCl2, 0.1 M
sodium cacodylate (pH 7.4)) for 30 min at 37 °C. The islets were centrifuged again
and the fixative was replaced with more fixative for 2 h at room temperature. Finally
the fixative was replaced again with fresh fixative and left overnight at 4 °C before
being resuspended in PBS.
2.3 Molecular biology
2.3.1 Genotyping
Ear clips were taken from mice using an ear puncher. DNA was extracted by addition
of 75 µl of alkaline lysis reagent (25 mM NaOH, 0.2 mM EDTA pH 8.0) to the ear
punches and incubating at 98 °C for 1 h. After this, neutralisation reagent (75 µl; 40
mM Tris HCl) was added prior to centrifugation at 14000 rpm 1 min. Extracted DNA
was ready to use in genotyping PCRs.
2.3.2 RNA extraction
Cells or islets were washed once in PBS followed by addition of 1ml TRIzol
(Invitrogen) for cells and 500 µl for islets in a 1.5 ml Eppendorf tube. After this 200
µl of chloroform (Sigma) per ml of TRIzol were added, the tube was vortexed and left
at room temp for 2-3 min until two layers formed. Samples were centrifuged at
12000 rpm for 15 min at 4 °C. The upper aqueous phase was removed and RNA was
precipitated by adding 400 µl isopropanol (Sigma) per ml of TRIzol used initially to
the retrieved aqueous phase. The samples were vortexed and incubated at room
temperature for 15 min before spinning for 15 min at 4 °C. The pellet was washed
twice in 1 ml 75% ethanol and centrifuged at 7500 rpm for 5 min. The dried pellet
69
was finally resuspended in nuclease-free water. The same quantity of RNA was used
from each sample to perform an RT-PCR and cDNA synthesis using random primers.
2.3.3 cDNA conversion
cDNA was generated from RNA using a High Capacity Reverse Transcription kit
(Applied Biosystem). A 1X reaction included 2 μl of each 10X RT Buffer, 10X RT
Random Primers, 0.8 μl 25X dNTP Mix (100mM), 1 μl Multiscribe Reverse
Transcriptase. The amount of H20 depended on the amount of RNA used. The
reaction was made up to 20 μl with the addition of nuclease-free H20. A maximum of
1μg RNA was converted to cDNA. For conversion the following protocol was carried
out in a thermocycler: 10 min 25 °C, 2 h 37 °C, 5 min 85 °C, hold 4 °C.
2.3.4 Real Time PCR
For gene expression measurements cDNA (2μl) from the RT-PCR reaction was used
as template for quantitative real time PCR using a Fast SYBR Green Master Mix
(Invitrogen). A 1X reaction was made up of 2 μl template, 6 μl SYBR Green, 0.35 μl
forward and 0.35 μl reverse primers, 3.3 μl H20. Each sample was amplified in
duplicates. The reaction was initiated at 50 °C for 2 min followed by the activation
and pre-denaturation step at 95 °C for 10 min. The run was made up of 40 cycles of
15 s at 95 °C and 1 min at 60 °C. mRNA levels were expressed as ΔCt and β-actin was
used as an endogenous control for normalisation.
2.3.5 Fluorescence activated cell sorting (FACS)
After overnight incubation, 250-300 islets of the same genotype were handpicked in
a 15 ml falcon and washed twice with 5 ml Hank’s based cell dissociation buffer
(Invitrogen) containing 0.1% BSA. They were then centrifuged at 2000 rpm for 2 min.
Islets were dissociated by repeated pipetting (80-100 times) in 150 μl of the solution
and 20 μl trypsin. The reaction was stopped with the addition of 20 μl FBS (Seralab).
The cells were washed in 1ml PBS, centrifuged at 2000 rpm for 1 min and
70
resuspended in 500 μl PBS and 1% FBS. 1 µg/ml DAPI (Roche) was added to stain for
dead cells. After a 10 min incubation the cells were filtered through a 35 μm cell
strainer. RFP+ and RFP- cells were recovered in 500 μl TRIzol after sorting on a BD
FACS ARIA III (BD Bioscience).
2.3.6 FACS analysis
Islets were dissociated into single cells as previously described, washed in PBS and
centrifuged at 600 g for 2 min. Cells were incubated in 50 μl 1:1000 near IR dead cell
stain (Life technologies) for 20 min at 4 °C, washed with PBA (PBS, 1% BSA, 0.1%
azide) and fixed in 2% PFA for 10 min at room temperature. They were then washed
twice with PBA and once with Saponin (0.025 % in PBA) before 10 min incubation
with Saponin at room temperature. Cells were incubated with primary antibodies
against ZnT8 (Mellitech), insulin (DAKO) and glucagon (Sigma) in 1:100 dilution in
Saponin for 20 min at room temperature. They were then washed twice in Saponin
before incubation with secondary antibodies (anti-mouse AF 405, anti-guinea pig AF
488, anti-rabbit AF 640) for 20 min at RT. Next were two final washes in Saponin
before resuspending in PBA. The samples were run on a BD Fortessa Flow
Cytometer (BD Bioscience).
2.4 Cloning and virus production
2.4.1 Competent bacteria
A trace of competent XL-1 Blue bacteria was taken with a sterile inoculating loop,
streaked out on a luria broth (LB) agar plate and incubated at 37 °C overnight. A
single colony was picked to inoculate 10 ml LB and incubated in a shaker at 37 °C
overnight. Overnight culture of 1 ml was added to pre-warmed LB and incubated on
the shaker at 37 °C until an appropriate density was reached (approximately 3.5 h).
The culture was cooled down on ice for 5 min before transfer into 50 ml centrifuge
tubes and centrifugation at 4000 g for 5 min at 4 °C. The supernatant was discarded
71
and cells were kept on ice. The following steps were performed in the cold room.
Cells were resuspended in 30 ml ice-cold TFB1 buffer (100 mM RbCl, 50 mM MnCl2,
30 mM KC2H4O2, 10 mM CaCl2, 15% glycerol) and kept on ice for 90 min. Cells were
collected by centrifugation at 4000 g for 5 min at 4 °C and then resuspended in 16 ml
ice cold TFB2 buffer (10 mM MOPS, 10 mM RbCl, 75 mM CaCl2, 15 % glycerol).
Aliquots (200 µl) were prepared in 1.5 ml Eppendorf tubes. These were frozen down
in a dry-ice ethanol bath. Competent cells were stored at -80 °C.
2.4.2 Transfection
Cells were seeded in 6-well plates and transfected the following day using
Lipofectamine-2000 (Invitrogen). Plasmid DNA (1 µg) and 3 µl Lipofectamine were
used per well. These were mixed separately into Optimem (100μl) and incubated at
room temperature for 5 min before mixing and further incubation for 20 min. 200 µl
of this solution and 1 ml Optimen were added to each well before culture overnight
at 37 °C in a humidified incubator. This was then replaced with normal culture
media. Imaging was done 48 h following transfection.
2.4.3 Bacterial transformation
Plasmid DNA (~10 ng) was added to 100µl of competent cells. The bacteria were left
on ice for 15 min before heat shock at 42°C for 1 min. Next, the cells were incubated
for 1h at 37 °C with constant shaking after addition of 300 µl of super optimal broth.
Cells (50μl) were plated on a warm LB/agar plate which included the suitable
antibiotic at a concentration of 100µg/ml.
2.5 Statistical analysis
Student’s t-tests were used to determine any significant difference between two
independent variables. Interactions between multiple treatments were examined
using two-way ANOVA (with adjustment for repeated measurements as necessary)
72
prior to pairwise comparisons with Bonferroni’s post hoc test. Analysis was
performed with GraphPad Prism 6.0. A value of P<0.05 was considered significant.
Data are expressed as means ± SEM.
73
2.6 Primers
Gene
Sequence
ZnT1 FOR: CAGGCAGAGCCAGAAAAATTG
REV: TGGATGAGATTCCCATTTACTTGTAC
ZnT2 FOR: CCCGACCAGCCACCAA
REV: CCAAGGATCTCGGCTCGAT
ZnT3 FOR: GGAGGTGGTTGGTGGGTATTT
REV: CAAGTGGGCGGCATCAGT
ZnT4 FOR: CATCGCTGCCGTCCTCTAC
REV: ATTTGCCATGTATCCACCTACAAG
ZnT5 FOR: ACTGTTTGCTGCCCTGATGAG
REV: CTCTATTCGGCCATACCCATAGG
ZnT6 FOR: ACGTCTCTGAAGCTGCTAGTACGA
REV: CCACACAAGCTGCGGCTAA
ZnT7 FOR: TGTGCCTGAACCTCTCTTTCG
REV: GCCTAGGCAGTTGCTCCAGAT
ZnT8 FOR: TGCACAGTCTACACATCTGGTCACT
REV: TGGCTGGCAGCTGTAGCA
ZnT9 FOR: GCACTGGGCATCAGCAAAT
REV: GAAAAGCCGTACGGGTGAGA
MT1 FOR: CCTTCTCCTCACTTACTCCGTAGC
REV: GGAGCCGCCGGTGGA
MT2 FOR: TCCTGTGCCTCCGATGGAT
REV: TGCAGGAAGTACATTTGCATTGT
MT3 FOR: AAGTGCAAGGGCTGCAAATG
REV: CACAGTCCTTGGCACACTTCTC
Actin FOR: CGAGTCGCGTCCACCC
REV: CATCCATGGCGAACTGGTG
74
Glucagon FOR: CCAAGAGGAACCGGAACAAC
REV: CCTTCAGCATGCCTCTCAAAT
Insulin FOR: CGTGGCTTCTTCTACACACCC
REV: AGCTCCAGTTGTGCCACTTGT
Abcc8-SUR1 FOR: GCCTACGCATCTCAGAAACCA
REV: CCATCTTGTACCTTTGCTTATTGAAG
Arx FOR: TGGGTCTGAGCACTTTTCTAGGA
REV: GAAAAGAGCCTGCCAAATGC
Gck FOR: CACAATGATCTCCTGCTACTATGAAGA
REV: AGCCGGTGCCCACAATC
Pcsk2 FOR: GAGTCCGAAAGCTCCCCTTT
REV: TTTGCAAGCCCATTGTGATAAA
Human ZnT8 FOR: CTGTCATCGAAGCCTCCCTC
REV: AAGGGCATGCACAAAAGCAG
Cre FOR: ATGTCCAATTTACTGACCG
REV: CGCCGCATAACCAGTGAAAC
ZnT8 flox FOR: AGTTATTGACTGAACACACCTATCTTA
REV: GCTATATACTCTTCCACTCAGCTACAT
RFP FOR: CTACAGGAACAGGTGGTGG
REV: CTGTTCCTGGGGCATGGC
Beta catenin FOR: AAGGTAGAGTGATGAAAGTTGTT
REV: CACCATGTCCTCTGTCTATTC
GlurtTA FOR: CATAAACGGCGCTCTGGAATTACTCAATGGAGTCG
REV: GCCGAGATGCACTTTAGCCCCGTCGCGATGTGAGA
Luciferase FOR: CAACTGCATAAGGCTATGAAGAGA
REV: ATTTGTATTCAGCCCATATCGTTT
75
2.7 Antibodies
Primary
Antibody Species Company Dilution
ZnT8 Rabbit Mellitech 1:200
Glucagon Mouse Sigma 1:1000
Glucagon Rabbit Santa Cruz 1:200
Insulin Guinea pig Santa Cruz 1:200
RFP Rabbit Clontech 1:100
Secondary
Antibody Species Company Dilution
Anti-mouse Alexa 568 Goat Invitrogen 1:1000
Anti-rabbit Alexa 488 Goat Invitrogen 1:1000
Anti-guinea pig Alexa
488 Donkey Invitrogen 1:1000
Anti-mouse Alexa 488 Donkey Invitrogen 1:1000
Anti-rabbit 568 Goat Invitrogen 1:1000
76
Chapter 3
ZnT8 deletion from a subpopulation of α-cells enhances
hypoglycaemia-induced glucagon secretion
77
3.1 Introduction
When the function of the hormone-producing α- and/or β-cells becomes impaired
this is likely to impact the regulation of glycaemia and could eventually cause
disease. These changes are associated with both T1D and T2D (see Chapter 1).
In addition to alterations in insulin release [289], glucagon release is also defective
and contributes to diabetic hyperglycaemia and impaired glucose homeostasis in
advanced stages of both disease types. In both cases, the α-cell no longer responds
normally to glucose concentrations so that the counter-regulatory response to
hypoglycaemia is weakened. Conversely, raised glucose levels, which should inhibit
glucagon secretion, elicit inappropriately high glucagon responses [290].
Genome-wide association studies examine SNPs across the genome and seek to
assess which, if any, may contribute to disease risk. Such research has so far led to
the discovery of ~90 loci in the human genome which appear to have a significant
impact on T2D risk [203, 205, 212, 291, 292]. Few of the SNPs have a known
functional impact on the amino acid sequence, with the majority of them located in
non-coding parts of the genome.
One of these loci harbours SLC30A8 which encodes the Zn2+ efflux transporter ZnT8.
The expression of this gene is largely restricted to the pancreatic islets where the
transporter resides on the membrane of the dense core secretory granule in α- and
β-cells [203].
A non-synonymous SNP rs13266634 results in an amino acid substitution from a
tryptophan to an arginine in the cytoplasmic C-terminus of the protein. This risk
variant has one of the strongest effects on T2D risk, 15% per inherited allele. The
transcript is believed to be translated into a protein with lower Zn2+ activity making
the transporter less efficient at Zn2+ accumulation across the membrane into the
granule [222, 259]
78
A considerable amount of research has been focused on endeavouring to take
information from GWAS towards eventual clinical exploitation by exploring the
function of the associated genes in vivo in mouse models. Various groups have
looked into the effect of modulating ZnT8 expression in the mouse by deleting the
gene either globally or specifically in the β-cell. Given ZnT8’s subcellular localization,
and the importance of Zn2+ in insulin biosynthesis and storage [267], the role of the
transporter has largely, if not almost completely, been studied in the context of the
β-cell and insulin secretion.
3.2 Aims
Although many research groups have done extensive work looking into the role of
ZnT8 in the β-cell, work on the α-cell is lacking and calls for further investigation.
Here we aimed to generate a mouse deleted selectively for ZnT8 in the α-cell and
investigate how this affects glucose tolerance and glucagon secretion in vivo.
79
3.3 Results
3.3.1 Expression of ZnT8 in the mouse α-cell
Dissociated islets from C57BL/6 mice were stained for glucagon and ZnT8 and
imaged by spinning disk confocal microscopy. ZnT8 immunoreactivity was present in
glucagon-positive cells and there was colocalization between the two stains.
However, the degree of colocalization was far from complete (Pearson correlation=
0.79) (Figure 3.1A). The presence of ZnT8 in the α-cell was also confirmed at the
mRNA level after extracting total RNA from fluorescence activated cell sorting (FACS)
purified α-cells. This revealed that ZnT8 is the most highly expressed member of the
ZnT family of transporters in these cells (Figure 3.1B)
Figure 3.1 – ZnT8 expression in the α-cell. (A) Dissociated islets cells were stained for
glucagon using anti-glucagon (Sigma, 1:1000, red) and anti-ZnT8 (Mellitech, 1:200, green) to
confirm the presence of ZnT8 at the protein level. Scale bar = 10 µm (B) Total RNA was
extracted from FACS-sorted cells to verify the expression of ZnT8 in the α-cell at the mRNA
level. Data from n=3 mice
Zn T1 Zn T2 Zn T3 Zn T4 Zn T5 Zn T6 Zn T7 Zn T8 Zn T9
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
-a
cti
n)
Glucagon ZnT8 A
B
80
3.3.2 Generation, breeding and phenotyping of mice null for ZnT8 in the α-cell
Animals bearing floxed ZnT8 alleles, originally obtained from GenOway (Lyon,
France) and backcrossed with C57BL/6 mice, were maintained on a standard chow
diet. The strategy followed by GenOway to generate ZnT8fl mice required the
insertion of LoxP sites, one upstream of exon 1 and a second one connected to a FRT
flanked neomycin selection cassette upstream of exon 1 and within intron 1 of the
Slc30a8 gene. The construct was homologously recombined in SV129-derived
embryonic stem cells at the ZnT8 site and was then injected into C57BL/6
blastocytes, before being transferred into pseudopregnant C57BL6 females. Crossing
with a flp-expressing strain then removed the neomycin cassette. The resulting
animals were further backcrossed 6 times to C57BL/6 mice.
The ZnT8fl/fl mice were crossed with mice carrying a Cre transgene under a 0.6 kB
fragment of the pre-proglucagon promoter (PPG) [280]. Further crossing was then
needed to generate homozygous mice for the floxed gene containing a single copy of
the transgene, rendering them ZnT8 KO (Figure 3.2A). The genotypes of the offspring
were identified by PCR analysis of genomic DNA extracted from ear biopsies. The
samples were tested for the presence of ZnT8 floxed alleles (Figure 3.2B) and Cre
transgene to determine the KO animals and those that would act as control (Figure
3.2C). This strain of mice was used in in vivo experiments and glucagon secretion
assays.
Morphologically α-cells are almost identical to the more abundant β-cells. The lack of
reliable and precise methods of distinguishing α-cells in the intact islet makes
research on α-cells challenging. Thus, this project also utilized transgenic mice
bearing a fluorescent protein for identification of α-cells in situ generating a triple
transgenic. ZnT8fl/fl mice with the Cre transgene under the control of the PPG
promoter were crossed with mice containing a stop-flox red fluorescent protein
variant (tandem dimer RFP, tdRFP/RFP) at the Rosa26 locus. In the presence of Cre
recombinase, recombination events flip the RFP construct from the antisense
Heterozygote
81
orientation into the sense. Another recombination event then takes place during
which the stop-flox cassette is removed allowing transcription driven from a CAG
promoter to read through into the properly-oriented tdRFP gene [293]. Using a Cre
maintained under the control of the PPG promoter prevented expression of RFP in
any other tissue or cell type other than α-cell [280]. The breeding strategy followed
can be seen in Figure 3.2B. In addition to this colony a separate one had to be bred
with mice WT for the ZnT8 locus but still bearing the PPGCre transgene and RFP for
use in in vitro experiments as controls (Figure 3.2C).
Figure 3.2 – Generation of αZnT8 KO mice and genotyping by PCR. (A) Schematic
representing the recombination event taking place to generate αZnT8 null mice in the
presence of Cre. (B) PCR genotyping for the floxed or WT allele. The band at ~ 330 bp is from
the WT allele while the larger band at ~ 460 bp is from the floxed allele of ZnT8. (C) PCR
genotyping for the Cre transgene to identify WT and KO animals. The band at ~ 330 bp
verifies the presence of Cre and the band at ~220 bp is that for β-catenin used as PCR
control.
200 bp 300 bp
Control KO
ZnT8 targeted allele
LoxP LoxP
Exon1 Exon2
LoxP
Exon2
Cre
A
B C
300 bp 500 bp
Homozygote
WT
Heterozygote
82
A C B
Figure 3.3 – Breeding strategy followed to generate αZnT8 KO mice and necessary controls
(A) for in vivo experiments that required littermate controls but not RFP, (B) for RFP-tagged
ZnT8 KO α-cells, (C) for RFP tagged α-cells with WT ZnT8 alleles.
3.3.3 Recombination efficiency and ZnT8 deletion
To assess the degree of recombination occurring in mice containing the Cre
transgene, initially whole islets from Cre+RFP+ mice were stained for RFP alongside
glucagon or insulin (Figure 3.4A). Imaging of these islets showed that only about 45%
of glucagon-positive cells expressed RFP. An almost negligible (2%) proportion of
insulin-positive cells was also labelled with RFP but this was most likely due to non-
specific background staining (Figure 3.4B). Additionally, islets positive for RFP from
WT and KO islets were stained for ZnT8 and glucagon or insulin to examine the
deletion of ZnT8 and not just the recombination efficiency. Immunohistochemical
analysis of the latter islets showed that ZnT8 immunoreactivity (Figure 3.5A) was
selectively eliminated from KO and not WT mice. However, only about half the
population of RFP+ cells showed a reduction in ZnT8 (Figure 3.5B). Insulin-positive
cells did not show a reduction.
83
W T K O
0
5 0
1 0 0
Pe
rce
nta
ge
(%
)
****
KO
WT
RFP ZnT8 A B
Figure 3.4 – Recombination in α-cells. (A) Representative images of PPGCreRFP+ islets were
stained for RFP (1:100) insulin (1:200) and glucagon (1:1000). (B) Images of stained islets
were used to calculate the percentage of glucagon and insulin positive cells expressing RFP.
Scale bar = 50µm **P<0.01, n=5-8 islets
Figure 3.5– ZnT8 deletion in α-cells. ZnT8 immunoreactivity (1:200) was assessed in WT and
KO islets expressing RFP. (A) Representative images showing ZnT8 in green and endogenous
RFP fluorescence in red. Scale bar = 10µm (B) The bar graph show the proportion of RFP+
cells that were also positive for ZnT8. n= 80-90 cells from 5-6 islets per genotype, Student’s
t-test, ****P<0.0001
To then get a more accurate quantification of the extent of deletion, isolated RFP+
islets from WT and KO mice were dissociated, fixed and stained for insulin, glucagon
and ZnT8 for FACS analysis (Figure 3.6A,B). This showed that approximately 30% of
glucagon-positive cells expressed RFP (Figure 3.6C), reflecting recombination at the
Rosa26 locus where the tdRFP reporter is located. Although clearly recombination in
G lu+
R F P+
In s+
R F P+
0
1 0
2 0
3 0
4 0
5 0
**
Pe
rce
nta
ge
(%
)
A B
84
these cells did occur at this locus, only 50% of RPF+ cells showed deletion of ZnT8
and thus recombination at the Slc30a8 locus. This suggests that Cre is more efficient
at recombination at the RFP than at the ZnT8 locus.
In addition, ZnT8 was found to be absent from a small percentage of glucagon-
positive RFP- cells indicating that recombination happened in a small proportion of α-
cells not expressing RFP (Figure 3.6D) Concomitantly, insulin-positive cells did not
show any changes in ZnT8 levels and they were all ZnT8-positive (Figure 3.6D).
Overall, ZnT8 expression was eliminated in ~15% α-cells (Figure 3.6D).
Such a low degree of deletion made us look into the use of other promoters as a
means of deleting ZnT8 in a greater proportion of α-cells. In the lab there was an
alternative strain that could have been used, with Cre again under a longer fragment
(>100 kbp) of the PPG promoter (iGluCre), present in a bacterial artificial
chromosome construct [294]. Islets from these iGluCre mice crossed with RFP mice
were extracted and stained to check for the level of recombination in this model. It is
evident from the images of islets stained for RFP and insulin (Figure 3.7) that RFP
expression is not restricted to α-cells since there is considerable double staining of
RFP and insulin making this delete strain unsuitable for use in this study.
3.3.4 Body weight and growth
αZnT8 KO mice were weighed regularly to check whether they had any variances in
weight with respect to control animals. Across all the ages examined, WT and KO
mice had a comparable body weight (Figure 3.8). Additionally, all the mice appeared
healthy with no obvious differences in growth.
85
Figure 3.6 – FACS analysis of WT and KO islets. Single islets following dissociation of RFP+
WT and KO islets were fixed, permeabilized and stained for insulin, glucagon and ZnT8. Cells
were stained prior fixation with a live/dead stain to only allow the inclusion of cells that
Zn T 8+ - c e lls Zn T 8
+ - c e lls
0
2 0
4 0
6 0
8 0
1 0 0
* * *
% o
f c
ell
s
R F P+
R F P+
Zn T 8+
R F P-
Zn T 8+
0
2 0
4 0
6 0
8 0
1 0 0 W T
K O
* * * *
% o
f c
ell
s
B
A
1) Single cells
2) Live
β-cells – RFP α-cells – RFP RFP+ α-cells – ZnT8
RFP_
α-cells – ZnT8
C D
1) Single cells
2) Live
β-cells – RFP
β-cells – ZnT8
α-cells – RFP
α-cells – ZnT8
RFP+ α-cells – ZnT8
RFP_
α-cells – ZnT8
β-cells – ZnT8 α-cells – ZnT8
86
were alive just before fixation. Representative traces showing expression analysis of cells
from WT (A) and KO (B) islets. The red traces denote the positive populations while blue
traces the negative populations. (C) Expression of RFP and ZnT8 from WT and KO islets as a
percentage of live cells. (D) ZnT8 expression in glucagon- and insulin-positive cells. Data are
mean ± SEM, n=3, two-way ANOVA, *P<0.01, ***P<0.0005.
3.3.5 Glucose homeostasis
In order to investigate the effect of deleting ZnT8 in α-cells on glucose homeostasis,
and particularly to detect whether these mice may display altered glucose tolerance,
we carried out intraperitoneal glucose tolerance tests (IPGTTs) on αZnT8 KO mice.
Littermates lacking the Cre transgene were used as controls to minimize variability
(Colony 1, Figure 3.3). It has been reported that age differences can have a huge
impact on glucose homeostasis in rodents [295] so IPGTTs were performed at 8, 12
and 16 weeks of age in case any effect might be age dependent. A glucose load of 1
g/kg was injected intraperitoneally and circulating glucose levels monitored at
various time intervals by taking blood from the tail. As can be seen in Figure 3.9
there was no discernible difference at any age examined.
Figure 3.7 – Immunostaining of iGluCre
islets. RFP+ islets bearing Cre under the
alternative longer PPG promoter [294]
were fixed and stained for insulin.
Endogenous RFP can be seen in red and
insulin in green.
insulin merge RFP
87
0 3 0 6 0 9 0 1 2 0
0
5
1 0
1 5W T
K O
T im e (m in )
Glu
co
se
(m
M)
0 3 0 6 0 9 0 1 2 0
0
5
1 0
1 5
T im e (m in )
Glu
co
se
(m
M)
W T
K O
0 3 0 6 0 9 0 1 2 0
0
5
1 0
1 5
T im e (m in )
Glu
co
se
(m
M)
W T
K O
A C B
Figure 3.8 – Weight measurements. The mice were weighed regularly shortly after they
were weaned up to the age of 16 weeks. There was no difference between WT and KO mice
either in females (A) or males (B).
Figure 3.9 – Glucose homeostasis. Intraperitoneal glucose tolerance tests were performed
on WT and αZnT8 KO mice fed a normal chow diet after an overnight fast at 8 (A) 12 (B) and
16 (C) weeks of age. n=8-15 mice/genotype
Another factor investigated was whether fasting conditions elicited a different
response from the KO mice. Blood glucose levels were measured after an overnight
fast and blood samples were taken from the tail vein for serum glucagon
measurements (Figure 3.10). Again no difference was observed in fasting blood
glucose levels or fasting glucagon compared to WT controls.
A g e (w e e k s )
We
igh
t (g
)
4 8 1 2 1 6
0
1 0
2 0
3 0W T
K O
A g e (w e e k s )
We
igh
t (g
)
4 8 1 2 1 6
0
1 0
2 0
3 0W T
K O
A B
88
Figure 3.10 – Fasting glucose and glucagon. The mice were fasted overnight and fasting
plasma glucose was measured using a glucometer (A). Plasma was taken for glucagon
measurements (B). n=7-8
3.3.6 Insulin sensitivity
In an effort to impose extra stress on the α-cell, and to test whether ZnT8 absence
affects insulin sensitivity, insulin tolerance tests (ITTs) were performed on mice of 8
weeks of age (Figure 3.11A). Blood glucose was measured at regular intervals and,
similarly to IPGTTs, there were no significant differences between WT and KO mice.
During the ITTs blood was collected at 0 and 15 min for plasma glucagon
measurements. The glucagon levels of WT and KO mice were very close at both time
points (Figure 3.11B).
Figure 3.11 – Insulin sensitivity. (A) Intraperitoneal insulin tolerance tests performed on 8
week old mice fasted for 5 hr and injected with 0.5U/kg insulin. (B) Plasma glucagon
measured at 0 and 15 min. n=13-14, *p<0.05
T im e (m in )
[glu
ca
go
n]
pg
/ml
0 1 5
0
5 0
1 0 0
1 5 0W T
K O*
*
0 1 5 3 0 4 5 6 0
0
5
1 0
1 5
T im e (m in )
Glu
co
se
(m
M)
W T
K O
A B
W T K O
0
2
4
6
8
Glu
co
se
(m
M)
W T K O
0
2 0
4 0
6 0
8 0
1 0 0
Glu
ca
go
n (
pg
/sa
mp
le)
A B
89
3.3.7 Streptozotocin treatment
In order to assess whether the deletion of ZnT8 might influence glucose homeostasis
in the context of diabetes, ZnT8 WT and KO mice at 6-8 weeks were rendered
diabetic by administration of streptozotocin (STZ), an agent which is selectively toxic
for the β-cell. STZ enters the cell through the GLUT2 transporter which is not
normally expressed in the α-cell [296]. The agent was administered to mice via the
intraperitoneal route for five consecutive days at a dose of 50 mg/kg. Glucose
homeostasis was assessed 4 weeks later by performing an IPGTT after an overnight
fast. The mice displayed fasting hyperglycaemia as expected though there were no
differences in blood glucose levels between WT and KO mice throughout the
duration of the test (Figure 3.12)
Figure 3.12 – Streptozotocin treatments.
Mice were injected with 50mg/kg
streptozotocin daily for 5 consecutive
days following overnight starvation.
IPGTTs were performed on WT and KO
mice 4 weeks later when the mice
showed signs of fasting hyperglycaemia.
n=8-9
3.3.8 Hypoglycaemic clamps
Since direct intraperitoneal insulin injections of insulin (Figure 3.11) could only be
used safely to produce relatively mild hypoglycaemia (4-5 mM) an alternative
approach was used to safely elicit a more marked reduction in blood glucose (2-3
mM) where glucagon secretion is likely to be more strongly stimulated. These
experiments were performed in Prof Christophe Magnan’s laboratory (University
Paris Diderot-CNRS UMR8251). To examine the in vivo response of these mice to low
glucose conditions the mice were rendered hypoglycaemic by infusion of insulin
(Figure 3.13A). Male mice did not exhibit any differences in the rate of glucose they
0 3 0 6 0 9 0 1 2 0
0
1 0
2 0
3 0
4 0W T
K O
T im e (m in )
Glu
co
se
(m
M)
90
0 9 0
0
2 0 0
4 0 0
6 0 0
8 0 0W T
K O
T im e (m in )
Pla
sm
a g
luc
ag
on
(p
g/m
l)
**
**
0 1 0 2 0 3 0 4 0 5 0 6 0 7 0 8 0 9 0
0
2
4
6
8
1 0
T im e (m in )
Gly
ca
em
ia (
mM
)
W T
K O
2 0 3 0 4 0 5 0 6 0 7 0 8 0 9 0
0
1 0
2 0
3 0
4 0
T im e (m in )
GIR
(m
g/m
in/k
g)
**
*
W T
K O
A
C
B
needed to maintain hypoglycaemia, however the female KO mice required a
significantly lower glucose infusion rate than littermate controls (Figure 3.13B). Such
a finding indicates a greater secretion of glucagon by KO mice. Serum glucagon
measurements performed on samples taken from these mice showed that at time
zero, KO mice had a lower basal release of the hormone but at 90 min after the mice
were rendered hypoglycaemic they released significantly more glucagon than WT
mice (Figure 3.13C).
Figure 3.13 – Hypoglycaemic hyperinsulinaemic clamps. These were performed on 12 week
old mice. They were rendered hypoglycaemic by administration of insulin (A). Glucose levels
were monitored and glucose infusion rate adjusted accordingly to keep the mice in a
hypoglycaemic state (B). (C) Plasma glucagon was measured from blood samples taken at 0
and 15 min. n=4-6. Data are mean ± SEM, two-way repeated measures ANOVA or two-way
ANOVA. *P<0.05, **P<0.01
91
3.4 Discussion
ZnT8 is a Zn2+ efflux transporter localized on the secretory granule membrane of β-
but also α-cells where it is believed to be involved in the storage of Zn2+ into the
granule from the cytoplasm [213]. In this part of the study we showed that ZnT8 is
not needed in the α-cell for normal glucose homeostasis or insulin sensitivity
although if a greater degree of deletion was achieved the findings could potentially
have been different. However, its absence from a subset of these cells causes an
exaggerated in vivo response in females under hypoglycaemic conditions imposed by
hypoglycaemic clamps.
It should be noted that mice bearing only the Cre transgene displayed normal
glycaemic control as assessed by fasting/refeeding studies and so were α-cell
responses to various regulators of secretion [297]. Similarly mice bearing only RFP
did not display any obvious defects and had normal fertility [293].
3.4.1 Recombination in the α-cell
FACS data from RFP+ dissociated islets stained for glucagon, insulin and ZnT8
revealed that the recombination efficiency of this mouse model is quite low. Thus,
based on the numbers of cells stained both for glucagon and ZnT8, there was only a
~15% reduction in ZnT8 expression in the α-cell population (Figure 3.6D), while there
was no significant deletion in the β-cell population. Thus although PPGCre appeared
to highly specifically delete ZnT8 in the α-cell and not β-cells or in other glucagon
expressing cells such as the gut or brain [280], this was only achieved in a minority of
α-cells. A low recombination rate has been described by others using Cre expressed
by the same PPG promoter. One report made similar observations to our own with
13% recombination [298], while others saw 35% [299] and 43% [300] deletion.
92
3.4.2 Glucose homeostasis and insulin sensitivity
Glucose tolerance tests assess the ability of the body to deal with glucose loads.
Given that ZnT8 was deleted only in the α-cell any differences in the KO animals can
reasonably be attributed to an altered capacity of the α-cell to deal with glucose.
Previous published work showed that the response of αZnT8 KO mice to glucose
during IPGTTs was similar to controls [272]. However, this was done only at one age
so testing the mice at a range of ages was deemed necessary to make sure no age-
related differences were missed out. Thus the mice were tested at the ages of 8, 12
and 16 weeks but no apparent differences were observed (Figure 3.9).
Fasting blood glucagon was similar in WT and KO which explains why there were no
differences in fasting blood glucose either (Figure 3.10).
Various groups have worked on mice with global deletions of ZnT8 and in terms of
glucose tolerance as assessed by IPGTTs. Two groups reported no change [268, 270]
while three mouse colonies were reported as glucose intolerant two of which only
displayed this effect at a specific age [222, 269]. Deleting ZnT8 in the β-cell caused
glucose intolerance in only one of the two studies [271, 272].
Insulin sensitivity assessed by insulin tolerance tests did not appear to be altered
either in αZnT8 KO mice (Figure 3.11). However, although blood glucagon levels
were similar, the blood glucose concentration did not go below the threshold
required for stimulation of glucagon secretion in either WT or KO mice. Thus what
we were seeing was not α-cells at their most active. This could be why no difference
was observed in this setting. By contrast in conditions of true hypoglycaemia
occurring during hyperinsulinaemic clamps the KOs behaved differently to the WTs.
Higher insulin concentrations were also tried in ITTs in order to increase the degree
of hypoglycaemia further. However this often resulted in a dramatic decrease of
glucose concentration which made necessary the administration of glucose or even
93
death as the glucose levels dipped rapidly. Thus these mice could not be included in
the results and a lower insulin level was deemed preferable.
3.4.3 Response to hypoglycaemia in vivo
Clamp studies have shown that female mice in a state of hypoglycaemia secrete 2-
fold more glucagon compared to control mice (Figure 3.13). It is interesting to note
that when ZnT8 was deleted in the β-cell, which functions during glucose
concentrations that are inhibitory to the α-cell, during stimulatory conditions the
opposite had been reported. That is insulin secretion in vivo is hindered by the
absence of ZnT8 [271]. Thus, deleting ZnT8 in these two cell types, which have
contrasting functions, causes an effect in the opposite direction.
3.5 Conclusion
Based on the findings of this study ZnT8 is not required in the α-cell for normal
glucose homeostasis. However this transporter is necessary in a subset of the α-cell
population normal responses to hypoglycaemia in vivo.
94
Chapter 4
ZnT8 is necessary in the α-cell for normal cytosolic and
granular Zn2+ homeostasis
95
4.1 Introduction
Zinc is one of the most important transition metals in the human body. In its ionic
form Zn2+ is an essential component of many proteins and is vital for a multitude of
cellular processes including cell division, transcription and signalling [301]. The Zn2+
content of the pancreas is one of the highest in the body [233] consistent with the
findings that these ions have a key role in insulin storage and secretion in β-cells
[266].
Diabetes has been known for many years to have an impact on Zn2+ homeostasis in
the body. Thus published work has shown that compared to healthy controls T2D
patients have a substantial reduction in total Zn2+ both in blood and at the cellular
level [236]. Similarly, murine genetic models of T2D exhibit lower pancreatic Zn2+
levels [238, 239].
Insulin in secretory granules is packaged in a hexameric structure co-ordinated by
two Zn2+ ions [267]. Correspondingly, autometallography has shown that Zn2+ in β-
cells is found in close association to secretory granules, with variable staining ranging
from heavy, sparsely or non-stained granules [302]. Although less intense the
staining was also present in α-cell secretory granules, mainly in the periphery, and all
α-cells examined under the electron microscope were stained to an extent [302].
Thus, the presence of Zn2+ in α-cell secretory granules implies that it could serve an
important function in the regulation of glucagon secretion and not just insulin
secretion. The fact that the Zn2+ transporter ZnT8 has been found to be encoded by a
T2D associated gene [203] further reinforces the likelihood of Zn2+ playing an
important role in islet cells. However it has generally been assumed that the role of
Zn2+, and thus of ZnT8 variants, in altering diabetes risk is mediated via alteration in
insulin release.
96
Many studies either on global or β-cell specific ZnT8 KO mice, have produced
controversial findings as to its role in the β-cell (see Introduction, chapter 1.13.1).
Although some groups have shown no differences in the majority of glycaemic and
other parameters tested, notably glucose tolerance and glucose-stimulated insulin
secretion, others have found that the absence of ZnT8 had a significant impact. This
included impaired glucose tolerance [222, 269], insulin secretion in vivo [222, 271],
reduced intracellular Zn2+ accumulation [222, 268] and morphological changes to
granule infrastructure [222, 270].
The α-cells of the pancreatic islet are electrically excitable, using electrical signals to
couple changes in plasma glucose levels to altered glucagon secretion, responding to
direct and indirect stimuli. Several candidates have been suggested to affect
glucagon secretion in a paracrine manner, insulin and other β-cell products such as
Zn2+ being some of these candidates. The cellular compartment with the highest
amount of Zn2+ in β-cells is the secretory granule, containing up to 70% of total Zn2+
and reaching a concentration of ~10-20 mM [129]. Upon exocytosis, due to the
increase in pH from ~5.5 in granules to ~7.4 in blood, the insulin crystals become
destabilized releasing both Zn2+ ions and creating a large local concentration of Zn2+.
Various studies have therefore raised the possibility for Zn2+ acting as a paracrine
regulator of glucagon secretion. Studies on rodent islets and cells have shown an
inhibitory effect of Zn2+ on glucagon release [121, 130].
Given the findings that ZnT8 appears to be important in the α-cell for the normal
regulation of glucagon secretion (Chapter 3) the impact of the absence of ZnT8 on
glucose control of signalling to both Zn2+ and Ca2+ in the α-cell was investigated.
97
4.2 Aims
Here we wanted to examine the effects of deleting ZnT8 in the α-cell at the single
cell level particularly to check whether this transporter is important in regulating
cellular Zn2+ homeostasis as observed in the β-cell. This was feasible using mice with
RFP-labelled α-cells for imaging live cells or identification for other in vitro
experiments.
98
4.3 Results
4.3.1 Impact of α-cell ZnT8 deletion on glucagon secretion in vitro
Mice lacking ZnT8 selectively in the α-cell were generated and maintained as
described in Chapter 3. The deletion of Slc30a8 specifically in the β-cell has
previously been shown to influence in vitro insulin secretion [252]. To determine
whether islets from αZnT8 KO mice respond differently to those from WT animals
glucagon secretion experiments were done on freshly isolated islets. WT and KO
islets were incubated with 1 mM or 17 mM glucose for 1 h and the glucagon released
was quantified using a radioimmunoassay. There was no change in secretion at 17
mM but at stimulatory conditions of 1 mM glucose, KO islets secreted significantly
more glucagon compared to controls (Figure 4.1).
Figure 4.1 - Glucagon secretion in vitro.
Isolated islets were incubated in 1 mM
and 17 mM glucose for 1hr. The
glucagon was the measured using an
immunoassay. The KO islets secreted
significantly more glucagon compared to
control. Data are mean ± SEM, n=5,
**P<0.01
4.3.2 Calcium responses
Differences in secretion capacity at the level of the individual α-cell might be caused
by altered intracellular free Ca2+ changes in response to low glucose concentrations
[303]. Thus, the impact of glucose on α-cell intracellular free Ca2+ levels was
investigated in whole islets by intracellular Ca2+ imaging with Fluo-2, an
intracellularly-trappable fluorescent Ca2+ indicator (Figure 4.2A). RFP+ WT and KO
islets were used in this instance for identification of α-cells and these were imaged
during exposure initially to 1mM then to 11 mM glucose (Figure 4.2B,C). The peak
1 1 7
0 .0 0
0 .0 5
0 .1 0
0 .1 5
0 .2 0
0 .2 5W T
K O
**
G lu c o s e (m M )
Glu
ca
go
n r
ele
as
e %
to
tal
99
Ca2+ levels apparent in the traces obtained (AUC; Figure 4.2D) and the area under
the curve (Figure 4.2E) showed that Ca2+ responses were not altered in the KO islets
compared to controls. Similarly, Fourier Transform analysis of the traces showed no
differences in the average frequency of oscillations (Figure 4.2F). Thus, exaggerated
Ca2+ responses to glucopenia cannot explain the increased glucagon secretion at 1
mM. Additionally we performed the statistical D’Agostino Pearson omnibus
normality test (Graph Pad Prism) which both WT and KO islets passed, showing that
the distribution of frequencies was Gaussian and there was no evidence for the
existence of two populations of cells.
Figure 4.2 – Low glucose-induced calcium responses. RFP+ WT and KO islets were incubated
with the calcium indicator Fluo2 (A) for 40 min before imaging at 1mM and 11mM glucose.
Scale bar 50 µm. Representative traces from WT (B) and KO (C) α-cells. (D) Area under the
curve (AUC). (E) Spike amplitude as a % of baseline. (F) Fast Fourier transform of calcium
traces. Data are mean ± SEM, n = 16-18 islets from 5 mice/genotype.
0.0
0
0.0
1
0.0
2
0.0
3
0.0
4
0.0
5
0.0
6
0.0
7
0.0
8
0
2 0
4 0
6 0
8 0
K O
W T
D o m in a n t fre q u e n c y (H z )
Pe
rce
nta
ge
ce
lls
(%
)
T im e (m in )
F/F
min
0 1 0 2 0
1 .2
1 .6
2 .0
2 .4
G 1 G 11
T im e (m in )
F/F
min
0 1 0 2 0
1 .2
1 .6
2 .0
2 .4
G 1 G 11
W T K O
0
2 0 0
4 0 0
6 0 0
8 0 0
1 0 0 0
AU
C (
AU
)
W T K O
1 0 0
1 2 0
1 4 0
1 6 0
1 8 0
Sp
ike
am
pli
tud
e R
FP
+v
e c
ell
s
(% o
f b
as
eli
ne
)
RFP Fluo-2 A C B
D E F
100
4.3.3 α-cell area
One possible driver of increased glucagon secretion would be an increase in
pancreatic α-cell mass, and/or a change in the proportion of α- versus β-cells. To
assess this possibility, pancreatic slices from the above mice were stained for
glucagon and insulin (Figure 4.3A) and imaged to determine the α- and β-cell area. In
KO mice (Figure 4.3B) there was a tendency for increased α-cell area and islets from
α-ZnT8 KO mice displayed a significant increase in α-to-β cell area ratio.
Figure 4.3 – Pancreatic histology. (A) Slices from fixed WT and KO pancreata were stained
for insulin (green) and glucagon (red). (B) The relative area is shown as α:β cell ratio. Scale
bar 50 µm. Data are mean ± SEM, n=5 mice, 8-10 slices per mouse, *P<0.05
4.3.4 Intracellular Zn2+ imaging
Given its role in the β-cell [222], where ZnT8 is involved in the transfer of Zn2+ from
the cytoplasm into the granule, we anticipated that a similar role in the α-cell would
mean that its absence could have an effect on cytoplasmic Zn2+ levels. In our lab Zn2+
imaging in islets is routinely done using a recombinant Zn2+ probe (eCALWY-4)
packaged in adenoviral particles [288]. Initially this probe was used in αTC1.9 cells, a
mouse α-cell line, to determine the free cytosolic Zn2+ concentration in the α-cell.
The cells were first imaged without additions to get a baseline and then after adding
TPEN, a membrane permeable chelator, followed by excess Zn2+ with the ionophore
pyrithione (Figure 4.4A). This calibration allows the determination of cytoplasmic
W T K O
0 .0 0
0 .0 5
0 .1 0
0 .1 5
0 .2 0
0 .2 5 *
:
-ce
ll r
ati
o
A B
Glucagon Insulin
WT KO
101
0 2 0 0 4 0 0 6 0 0 8 0 0
3 .5
4 .0
4 .5
5 .0
5 .5
T P E NZ nP yr
T im e (s e c )
Cit
rin
e/C
eru
lea
n
6 0 0
7 0 0
8 0 0
9 0 0
1 0 0 0
[Zn
2+] c
yt
(pM
)
A B
[Zn2+] for each individual cell using the changes in FRET ratio which occur after the
aforementioned additions. Results from imaging with this probe (Figure 4.4B) on a
mouse α-cell line showed that cytoplasmic [Zn2+] in mouse α-cells ranged from 680-
920 pM averaging ~810 pM.
Figure 4.4 – Zn2+ imaging with eCALWY-4. αTC1.9 cells were infected with eCALWY-4
adenovirus bearing the cytosolic form of the probe. Cells were imaged the next day first at
baseline followed by the additions indicated on the representative trace (A). (B) Cytosolic
[Zn2+] values from individual α-cells.
Adenoviruses are very efficient at infecting primary β-cells but appear to be very
inefficient in the case of α-cells. This has been seen in our laboratory [304] and by
others [305] but was also observed by us after staining of fixed islets for glucagon
following infection of whole islets with eCALWY-4. Normally a multiplicity of
infection (MOI) of 20-25 [306] is used in experiments of this sort. In an effort to
assess whether a higher number of viral particles could be effective in driving
infection of α-cells, MOIs of 500, 2500 and 5000 were used for infection of islets
prior to fixation. Imaging of islets infected in this way showed that although there
was good infection of the islet as a whole, this only represented the β-cell population
since the glucagon-positive cells were still largely negative for the GFP-containing
virus (Figure 4.5). Increasing the MOI by 20 or even 200 fold did not appear to
detectably improve the infection of α-cells (Figure 4.5A-C).
102
Figure 4.5 – Islet expression of adenoviral Zn2+ probe. Islets infected with eCALWY-4
adenovirus were fixed and stained for glucagon. MOIs of 500 (A), 2500 (B) and 5000 (C) were
used to check if an increased viral load affected the efficiency of infection of α-cells.
In an effort to overcome this issue of very low infectivity by adenoviruses we sub-
cloned the virus into an adeno-associated virus (AAV) potentially allowing uptake by
a distinct set of cell surface receptors. In this case the viral genome had a flex switch,
so the coding sequence was placed in the opposite orientation to that required for
transcription. In the presence of Cre, which would recognize the lox sites, this
orientation would be flipped allowing expression of eCALWY4 (Figure 4.6A). This
would have been ideal for expression of the probe in the primary α-cells in our
mouse model which express Cre.
Positive clones were identified (Figure 4.6B) and AD293 cells were co-transfected
with a Cre-expressing plasmid showing robust expression of eCALWY-4, confirming
eCALWY-4 glucagon merge A
C
B
103
eCALWY-4 2.2kb
2Kb
A
C B
that the probe was functional in the AAV construct (Figure 4.6C). The virus was then
generated and tested but we could not detect expression of the probe in islets
positive for Cre in the α-cells nor in β-cells, even when using other AAVs containing
other recombinant proteins which were shown to work when used in the brain. It
thus seems that the AAV serotypes 1 and 5 harboured by the virus bearing the probe
are not effective at infecting α-cells or β-cells thus making this virus unusable in
primary islet cells.
Figure 4.6 – AAV-eCALWY-4. (A) DNA construct and recombination events for Cre-
dependent eCALWY-4 expression. (B) Double RE digest (SpeI, EcoRI) of positive colonies
containing the construct. Lanes 1, 3, 5-undigested DNA. Lanes 2, 4, 6-double digested DNA.
(C) Image of AD293 cells expressing eCALWY-4 following transfection with a Cre-pCDNA3.1
and eCALWY-4-pAAV plasmids
Since the adenoviral or AAV recombinant probe could not be utilised an alternative
was found, a loadable Zn2+ indicator FluoZin-3, which is expected primarily (though
perhaps not exclusively) to be localized to the cytoplasm [307]. RFP+ WT and KO
islets were incubated with the dye for an hour before imaging (Figure 4.7A) initially
perfusing with no additions to allow a recording of baseline fluorescence. At about
10 min the Zn2+ chelator TPEN was added, binding essentially all intracellular labile
Zn2+ and causing a reduction in fluorescence of the dye (Figure 4.7B). The
104
fluorescence value at the baseline and the minimum value following the addition of
TPEN were used to give a ratio which gave an indication of the free Zn2+
concentration in the cytoplasm. Note that the maximum fluorescence in the
presence of high intracellular Zn2+ concentrations after the addition of ZnCl2 to the
cells, in the presence of the ionophore pyrithione could not be used to provide a
precise calibration of fluorescence in terms of free Zn2+ it caused a saturation of the
signal (not shown). Nevertheless, based on the values determined as above,
intracellular Zn2+ levels appeared to be lower in KO compared to WT α-cells (Figure
4.7C). A similar difference was observed previously in whole islets from β-cell specific
KO animals, in this case using the recombinant probe eCALWY4 [222]. No such
difference was evident in non-RFP labelled cells which were largely made up of the
β-cell population of the islet, and in which ZnT8 was still expressed (Figure 4.7D).
Figure 4.7 – Intracellular Zn2+ imaging. RFP+ WT and KO islets incubated with the Zn2+
indicator FluoZin-3 for 1 h (A) were imaged at baseline (F1) and following perfusion with the
Zn2+ chelator TPEN (Fmin). (B) Representative trace from an α-cell. (C) Zn2+ levels of RFP+ cells,
quantified from the relative fluorescence (F1/Fmin). (D) Zn2+ levels of RFP- cells. Data are mean
± SEM, n = 16-20 islets from four mice, Student’s t-test, ***P<0.0001
W T K O
1 .0
1 .5
2 .0
2 .5
F1
/Fm
i (A
U)
0 1 0 2 0 3 0
3 0 0 0
4 0 0 0
5 0 0 0
6 0 0 0
7 0 0 0
8 0 0 0
T P E NF 1
F m in
T im e (m in )
Flu
ore
se
nc
e (
AU
)
W T K O
1 .0
1 .5
2 .0
F1
/Fm
in (
AU
)
****
RFP FluoZin-3
A
C
B
D
105
4.3.5 Impact of α-cell ZnT8 deletion on secretory granule Zn2+ content
Considering the proposed location of ZnT8 on secretory granules and its implied role
in maintaining granular Zn2+ concentration the next step was to search for any
changes in granular Zn2+ content. For this the RFP+ WT and KO islets were incubated
with the loadable Zn2+ dye Zinpyr4 [283]. Experiments were first done to check that
it colocalized with the granules. INS-1 (832/13) cells were loaded simultaneously
with the dye and a fluorescent Lysotracker [308] with an orthogonal emission
spectrum. The latter, although usually specific for lysosomes in non-secretory cells,
also enters insulin granules due to their mildly acidic environment and positive
charge with respect to the cytosol [309]. As shown in Figure 4.8A the staining not
only appeared granular but there was also good colocalization between Zinpyr4 and
the lysotracker, verifying the localization of the Zn2+ probe to the granule interior.
However, this dye photobleached very quickly even at the low laser powers used so
a decrease in signal was evident in real time when attempting to make a long
acquisition. It was thus not possible to acquire a similar trace as the one obtained
with the cytoplasmic dye FluoZin-3 (Figure 4.7B). Therefore, imaging of Zinpyr4 was
restricted to the acquisition of static images (Figure 4.8B) whilst all the conditions
such as exposure time, laser power and incubation time were kept the same to make
sure any variance in fluorescence was not a result of these variables being different.
The fluorescence intensity was measured using the corrected total cell fluorescence
equation (CTCF), a parameter which eliminates any discrepancy in actual
fluorescence due to cell size. Compared to RFP+ cells from WT islets, KO cells
exhibited an approximated 50% reduction in apparent free granular Zn2+ (Figure
4.8C).
4.3.6 Gene expression analysis
To confirm the deletion of ZnT8 at the mRNA level and to establish whether there
were any changes in the expression of other genes in the ZnT family, WT and KO
islets were isolated, dissociated and then sorted through a flow cytometer based on
RFP fluorescence. RNA was extracted from the RFP+ α-cells collected during the sort
106
and expression levels of ZnTs and MTs were assessed by qPCR. As previously shown
by FACS ZnT8 was not completely absent from the RFP+ population, with expression
in KO cells reaching about 40% that of WT cells (Figure 4.9A). Purified α-cells
expressed low levels of ZnT1, 4-7 and 9 while ZnT2 and 3 were not detectable (Figure
4.9A). Deleting ZnT8 caused no major changes in the expression levels of these
genes. WT cells also expressed the cytoplasmic Zn2+ binding proteins MT1 and MT2
but not MT3 with levels being comparable in WT and KO mice (Figure 4.9B). We also
looked into the expression levels of KATP channel subunits and Gck, in order to
determine whether there were changes in the energy sensing components of the
cell. Likewise we examined the expression of Arx and Pcsk2, whose gene products
are involved in α-cell identity and glucagon processing respectively (Figure 4.9C).
There were no changes in the expression levels of these genes nor in the expression
of glucagon in ZnT8 null α-cells (Figure 4.9D).
Figure 4.8 – Granular free Zn2+ imaging. (A) INS1 cells were incubated with ZinPyr-4 and
Lysotracker to determine the localization of ZinPyr-4 in secretory granules. (B) RFP+ WT and
KO islets were incubated with the Zn2+ dye Zinpyr-4 for 30 min before imaging Quantification
of fluorescence is shown as CTCF (Corrected Total Cell Fluorescence) (C) calculated using the
area of the cell chosen, integrated density and background fluorescence. Scale bar 10 μm.
Data are mean ± SEM, n=15-20 islets from three mice, Student’s t-test, **<0.002
W T K O
0
4 .01 0 5
8 .01 0 5
1 .21 0 6
CT
CF
**
RFP Zinpyr-4
KO
WT
Lysotracker Zinpyr-4 A
C B
D
107
Figure 4.9 – Gene expression analysis. RFP+ WT and KO islets were dissociated and then
sorted. The FACS sorted RFP+ cells were used for RNA extraction and qPCR. (A) Expression
levels of the ZnT family of transporters, (B) metallothioneins, (C) energy sensing and α-cell
specific genes, (D) glucagon in WT and KO cells. n= 5, ***P<0.001
4.3.7 Glucagon content
Increased glucagon content in KO α-cells might explain the enhanced glucagon
secretion observed at stimulatory glucose conditions. The possibility was examined
using two ways. One was to isolate pancreata from WT and KO mice and extract
proteins for total glucagon measurement per unit tissue with an HTFR assay. At the
pancreatic level no differences were observed in the glucagon content between WT
and KO mice (Figure 4.10C). The second method was to isolate RFP+ islets and stain
for glucagon following fixation. Then, the glucagon content of the RFP+ cells was
imaged by taking z-stacks which were then used to measure the total intracellular
glucagon volume and thus the content (Figure 4.10A). Again there were no
differences in total content comparing WT to KO cells (Figure 4.10B).
M T 1 M T 2 M T 3
0 .0
0 .1
0 .2
0 .3
0 .4
0 .5
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
ac
tin
)
W T K O
0
5 0
1 0 0
1 5 0
2 0 0
2 5 0
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
ac
tin
)
n s
A rx A b c c 8 K c n j1 1 G c k P c s k 2
0
1
2
3
4
W T
K O
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
ac
tin
)
Z n T 1 Z n T 2 Z n T 3 Z n T 4 Z n T 5 Z n T 6 Z n T 7 Z n T 8 Z n T 9
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
W T
K O
* * * *
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
ac
tin
)
A
C
B
D
108
Figure 4.10 – Glucagon content measurements. (A) WT and (B) αZnT8 KO islets were
stained for glucagon and imaged on a confocal microscope at 63x magnification. (C) RFP+
cells were selected for z-stacks and their glucagon volume was measured using ImageJ.
n=45-50 cells from 3 animals. (D) Pancreata from WT and KO mice were isolated and protein
was extracted using an acidic ethanol method and glucagon measured using an HTRF assay.
n= 3
W T K O
0 .0
0 .5
1 .0
1 .5
2 .0
Glu
ca
go
n c
on
ten
t
(n
g/m
g p
an
cre
as
)
W T K O
0
2 0
4 0
6 0
8 0
Glu
ca
go
n c
on
ten
t (
m3
)
A B
C D
109
4.4 Discussion
In this chapter we investigated the effect of ZnT8’s absence on α-cell in vitro and at
the single cell level. Work on isolated islets confirmed the finding of enhanced
glucagon secretion during hypoglycaemia (Figure 4.1). Zn2+ imaging showed a
reduction in free Zn2+ levels both intracellularly and in the secretory granules.
Additionally, we showed that the increased glucagon secretion observed during
hypoglycaemia both in vivo and in vitro cannot be explained by increased glucagon
content or by heightened Ca2+ responses to low (stimulatory) glucose concentrations
in the KO α-cell.
4.4.1 Glucagon secretion
Glucagon release from isolated islets at 1mM glucose was ~80% higher from KO
islets compared to WT controls (Figure 4.1). This is in agreement with the in vivo
findings of enhanced glucagon secretion in hypoglycaemic conditions (Figure 3.13).
Imaging stained pancreatic slices showed an increase in the α:β-cell ratio by ~15%
which could partly contribute to the increased glucagon secretion at low glucose.
However changes in some other cellular mechanisms must be taking place in the KO
mice that could explain this large increase in glucagon release.
Zn2+ homeostasis in the α-cell
ZnT8 was found to be important in Zn2+ homeostasis in the β-cell [222] and thus we
wished to investigate whether these ions played a similar role in the α-cell. First, we
examined cytosolic Zn2+ levels using a protein-based Zn2+ indicator. The recombinant
Zn2+ probe would have been preferable since it would be strictly targeted to the
cytosol and also it would allow us to use the ratio of fluorescence from the two
fluorophores it contained. Calibrations based on its dissociation constant for Zn2+
would then have allowed us to obtain values in molar terms for the free
concentration of Zn2+. Unfortunately, however, the traces obtained with the
110
alternative FluoZin-3 probe could not be calibrated accurately due to the dynamic
range of the indicator which meant that the maximum fluorescence was not
detected properly (i.e. the signal was saturated). Consequently these could not be
used in a calculation for Zn2+ concentration.
Despite this limitation we were able to use the FluoZin-3 fluorescence values at
baseline, and after the cell was depleted of Zn2+, to give a ratio which suggested that
Zn2+ levels in the cytosol were reduced in KO vs WT cells. Such a finding although
consistent with observations from the β-cell [310], are nonetheless somewhat
surprising. The lack of an efflux transporter is not expected to have resulted in a
reduction in cytosolic Zn2+. In fact, the change should have been in the opposite
direction. This indicates that ZnT8 may in normal conditions be acting as an influx
transporter at sites other than the granules such as the plasma membrane. Similar
findings have been made for another member of the ZnT family, ZnT5 [311].
Similarly, using ZinPyr-4 we imaged granular Zn2+ content. In this case the findings
were consistent with the proposed function of ZnT8 in transporting Zn2+ across the
granule membrane and into the granule lumen from the cytosol. As expected the
absence of ZnT8 led to a reduction in granular content (Figure 4.8C) reflecting a loss
of transporting activity in the KO cells.
4.4.2 Absence of effects of ZnT8 inactivation in α-cells on glucose-regulated Ca2+
fluxes
Interestingly, the absence of ZnT8, and consequent changes in cytosolic and granule
Zn2+, exerted no apparent effects on Ca2+ dynamics in α-cells, despite reports of
multiple interactions between these two ions on Ca2+, K+ and other channels [312].
Further, more detailed investigations possibly involving electrophysiology may be
needed to explore this surprising result in the future.
111
4.5 Conclusion
ZnT8 appears to be important in Zn2+ homeostasis in the α-cell influencing both
cytosolic and granular compartments leading to a reduction in concentration which
cannot be explained by changes in any of the other zinc transporters examined. This
direction of change in cytosolic concentration points towards a role for ZnT8 in Zn2+
influx and not just efflux. A mechanism other than enhanced calcium responses must
be responsible in causing the enhanced glucagon secretion during hypoglycaemia
possibly involving Zn2+.
112
Chapter 5
Overexpression of human ZnT8 in the murine pancreatic
α-cell leads to reduced glucagon release and impaired
responses to hypoglycaemia
113
5.1 Introduction
Functional studies in model systems are vital if we are fully to understand the
physiological role(s) of genes identified through genome-wide association and other
studies as affecting the risk of T2D [212]. Most studies up to now have used gene
deletion globally or in disease-relevant tissues to explore this question (see
Introduction, and Chapters 3,4) [210, 212]. Although investigating the absence of a
gene can be very informative its overexpression can likewise give similarly important
findings into the role of a protein. In the case of SLC30A8/ZnT8 this is of particular
importance since the impact of risk variants on activity and diabetes risk is still open
to discussion: the common risk variant (R325) is thought to lower activity and
increase risk [222], whereas rare truncated variants appear to be protective [221].
Inducible expression systems are commonly used in rodent systems to achieve both
temporal and spatial (i.e. tissue-specific) expression of a given gene. Components of
the Tet Switches [313] originate from the tetracycline (Tet) resistance operon in
E.coli belonging to one of the most evolved gene regulation systems. The Tet-Off and
Tet-On systems are used in the majority of the studies involving inducible
expression. The Tet-Off system was the initial system developed in 1992 and in the
presence of the antibiotic tetracycline the expression from a tet-inducible promoter
is decreased.
In order to utilize tetracycline as a regulator of transcription, a tetracycline-
controlled transactivator (tTA) was made by fusion of the tetracycline repressor with
a transcriptional activation domain from HSV. Thus, in the absence of tetracycline
the fusion protein can bind tet operator sequences and promote transcription while
in the presence of the antibiotic, its binding to the protein makes it unable to bind
DNA leading to a decrease in gene expression. The Tet-On system was later
developed by mutation of the repressor portion of the tTA so that it would rely on
114
tetracycline for induction of gene expression and not repression. This is known as a
reverse tetracycline controlled transactivator (rtTA) [314].
The system was first used in the β-cell [315] and about 10 years later the system was
utilised in the α-cell [316]. Recently, our laboratory has used this approach to
determine the effects of ZnT8 over-expression in the pancreatic β-cell in mice by
driving rtTA expression with the rat insulin promoter [317]. In the present study, the
rtTA sequence was placed under the control of the PPG promoter to generate
GlurtTA mice and to drive expression of this gene selectively in the α-cell in the adult
mouse.
5.2 Aims
Here, we wanted to investigate the effect the overexpression of ZnT8 would have on
glucagon secretion, GlurtTA mice were therefore crossed to mice bearing a human
ZnT8 transgene whose expression was driven by a tet operator sequence.
115
5.3 Results
5.3.1 Generation and genotyping of αZnT8Tg mice
For this project, GlurtTA mice were obtained (Dr Ahmed Mansouri, Max-Planck
Institute, Goettingen, Germany) and crossed to animals containing a human ZnT8-
overexpressing transgene (ZnT8Tg) as described [317]. Heterozygote GlurtTA mice
were crossed to homozygote ZnT8Tg animals. Two founder were used,
corresponding to lines #31 and #23 in Mitchell et al [317]. The litters comprised
GlurtTA offspring, which expressed the human ZnT8 transgene after induction with
doxycycline (as described above), while the littermates bearing the ZnT8Tg allele
alone were used as controls.
GlurtTA mice possess a 1.1kb region of the glucagon promoter upstream of the rtTA
[316]. In the α-cell the transactivator is produced but is not in its active form.
However doxycycline, a tetracycline derivative, binds and activates it so that it can
then in turn interact with the Tet operator sequence upstream of the construct
containing the cDNA of human ZnT8 and transactivate its expression. Transcription is
only possible as long as doxycycline is present. Consequently, the drug needs to be
administered continuously [316]. Mice were given doxycycline in the drinking water
(2g/L) from the age of 6 weeks.
The genotyping for GlurtTA was performed using standard PCR on DNA extracted
from ear biopsies (Figure 5.1). For the ZnT8 transgene, qPCR was used to detect the
presence in the genome of the luciferase gene, also included in the transgene.
Figure 5.1 – Genotyping by PCR. The
GlurtTA transgene was amplified by
standard PCR to identify control and
transgenic animals. Lanes 1, 2, 4, 6, 7, 9,
11 contain samples from transgenic
animals.
200 bp
Control
GlurtTA
1 2 3 4 5 6 7 8 9 10 11
116
5.3.2 Expression of human ZnT8 in the islet
Initially to check that the administration of doxycycline was effective in inducing the
overexpression of ZnT8, total RNA was extracted from isolated islets incubated
overnight with doxycycline (5µg/ml). qPCR analysis revealed expression of the
human form of ZnT8 in the mice containing both the overexpressing transgene and
the GlurtTA but not in the control mice carrying only the ZnT8 transgene (Figure 5.2).
We tested both founders that contained the transgene which showed that only
founder 2 (#31 from Mitchell et al [317]) expressed detectable levels of human ZnT8.
Consequently, this founder was used in all experiments.
Figure 5.2 – Human ZnT8
expression in isolated islets.
Total RNA was extracted from
islets from control and
transgenic mice. The levels of
hZnT8 were assessed by RT-
qPCR.
In order to obtain a more accurate assessment of the degree of overexpression in a
pure α-cell population and as opposed to islets, which are largely composed of β-
cells [318], isolated islets were dissociated, fixed and stained with glucagon for
fluorescent –activated cell sorting. RNA was then extracted from the sorted cells to
check for expression of the human ZnT8 in the α-cells obtained from the transgenic
and control animals (Figure 5.3).
c o n tro l fo u n d e r 1 fo u n d e r 2
0 .0 0 0
0 .0 0 1
0 .0 0 2
0 .0 0 3
mR
NA
ex
pre
ss
ion
(re
lati
ve
to
-ac
tin
)
117
Figure 5.3 – Hum an ZnT8 expression in
purified α-cells. RNA was extracted from
α-cells obtained from control or αZnT8Tg
mice. The levels of hZnT8 mRNA were
assessed by RT-qPCR. n=3
5.3.3 Intraperitoneal glucose tolerance was unaltered in α-ZnT8Tg mice
To check glucose homeostasis in these mice, IPGTTs were performed at the age of 8
weeks (Figure 5.4A) and 12 weeks (Figure 5.4B). Similarly to our findings from αZnT8
KO animals (Chapter 3) there was no difference in blood glucose between control
and transgenic animals at any time point during the test.
Figure 5.4 – Glucose tolerance in αZnT8Tg mice. Intraperitoneal glucose tolerance tests
were performed on control and transgenic mice at 8 (A) and 12 weeks of age (B). n=8-10
mice/genotype
0 3 0 6 0 9 0 1 2 0
0
5
1 0
1 5
Glu
co
se
(m
M)
C o n tro l
T ra n s g e n ic
0 3 0 6 0 9 0 1 2 0
0
5
1 0
1 5
T im e (m in )
Glu
co
se
(m
M)
C o n tro l
T ra n s g e n ic
A B
C o n tro l T ra n s g e n ic
0 .0
0 .2
0 .4
0 .6
0 .8
hZ
nT
8 m
RN
A e
xp
res
sio
n
(re
lati
ve
to
-ac
tin
)
118
5.3.4 Intraperitoneal insulin tolerance tests (ITTs)
ITTs were performed on transgenic and control male (Figure 5.5A) and female mice
(Figure 5.5B) at the age of 14 weeks in order to check their insulin sensitivity. The
responses appeared to be the same in the ZnT8 overexpressing and control mice.
5.3.5 Hypoglycaemic clamps
As discussed in Chapter 3, the above experiments could only be used safely to
provide a limited decrease in blood glucose levels (to ~4mM). In order to produce a
more marked decrease in blood glucose levels, where glucagon release is likely to be
more strongly stimulated hypoglycaemic clamps were performed on male mice in
Prof Christophe Magnan’s laboratory (University Paris Diderot-CNRS UMR8251). To
investigate the in vivo response to the imposed low glucose levels the mice were
rendered hypoglycaemic by continuous intravenous administration of insulin (Figure
5.6A) (see Methods, Chapter 2). ZnT8 overexpressing mice required a significantly
higher glucose infusion rate to maintain hypoglycaemia compared to control mice
(Figure 5.6B), suggesting an impairment in glucagon release. Glucagon levels were
consequently measured prior to the onset of hypoglycaemia and at 120 min after the
induction of hypoglycaemia (Figure 5.6C). This revealed lower plasma glucagon levels
at the latter time point, consistent with the higher glucose infusion rates required
during the clamps.
119
Figure 5.5 – Insulin sensitivity. Intraperitoneal insulin tolerance tests performed on 12
week old male (A) and female (B) mice fasted for 5 hr and injected with 0.5U/kg insulin. n=9-
10 mice per genotype
5.3.6 Glucagon secretion is impaired in islets isolated from αZnT8Tg mice
We sought next to determine whether the impaired glucagon secretion observed in
vivo in αZnT8 mice chiefly reflects a cell autonomous effect in the α-cell, as opposed
to a more complex set of changes in the autocrine response to hypoglycaemia,
potentially involving multiple tissues (brain, adrenals etc) [319]. Furthermore, we
have previously shown that deleting ZnT8 from the α-cell enhances glucagon
secretion from isolated islets (Chapter 3) and were therefore interested in the
hypothesis that overexpression may have an effect in the opposite direction i.e. to
impair glucagon secretion in response to low glucose. Correspondingly, we observed
that when isolated islets were incubated in 1 mM glucose (a concentration in the
range which strongly stimulates glucagon secretion from α-cells) [88], those
overexpressing ZnT8 secreted significantly less glucagon that control islets (Figure
5.7). At 17 mM glucose the responses did not differ between islets overexpressing
ZnT8 selectively in the α-cell and control islets.
0 1 5 3 0 4 5 6 0
0
2
4
6
8
1 0
T im e (m in )
Glu
co
se
(m
M)
C o n tro l
T ra n s g e n ic
0 1 5 3 0 4 5 6 0
0
5
1 0
1 5
T im e (m in )
Glu
co
se
(m
M)
C o n tro l
T ra n s g e n ic
A B
120
Figure 5.6 – Elevated glucose infusion rates in αZnT8 KO mice during hypoglycaemic
hyperinsulinaemic clamps. Experiments were performed on 12 week old male mice. Animals
were rendered hypoglycaemic by infusion of insulin (A). Glucose levels were monitored and
glucose infusion rate adjusted accordingly to keep the mice in a hypoglycaemic state (B). (C)
Blood glucagon levels were measured at 0 and 120 min. n=4. Data are mean ± SEM, *P<0.05,
**P<0.01, ***P<0.005.
Figure 5.7 - Glucagon secretion from αZnT8Tg
islets is impaired in vitro. Isolated islets were
incubated at 1 mM or 17 mM glucose for 1 h.
Released glucagon was measured using an
HTRF assay. Transgenic islets secreted
significantly less glucagon compared to control
when incubated at 1 mM glucose. Data are
mean ± SEM, n=4 separate experiments,
*P<0.05
1 1 7
0 .0 0
0 .0 5
0 .1 0
0 .1 5
0 .2 0
0 .2 5
C o n tro l
T ra n s g e n ic
G lu c o s e (m M )
Glu
ca
go
n r
ele
as
e %
to
tal *
0 2 0 4 0 6 0 8 0 1 0 0 1 2 0 1 4 0
0
5
1 0
1 5
T im e (m in )
Gly
ca
em
ia (
mM
)
C o n tro l
T ra n s g e n ic
0 2 0 4 0 6 0 8 0 1 0 0 1 2 0 1 4 0
0
2 0
4 0
6 0
T im e (m in )
GIR
(m
g/m
in/k
g)
C o n tro l
T ra n s g e n ic
***
***
0 1200
20
40
60
80Control
Transgenic **
Time (min)
Glu
ca
go
n (p
M)
A B
C
121
Efforts were next made to detect human ZnT8 by immunostaining using FACS
analysis and confocal microscopy. The antibody used (Mellitech) was described as
specific to human ZnT8. However, analysis of samples by both methods showed that
this might not have been the case. Thus staining of islets did not allow the detection
of human ZnT8 positive populations since there was high background (results not
shown). Similarly when using the antibody to try to identify human ZnT8-positive α-
cells by FACS the entire population of islet cells was found to be positive in both
ZnT8 overexpressing and control mice (results not shown). This was the case each
time regardless of titrating the antibody and using different dilutions to determine if
this was just a result of saturation. Thus, the antibody very likely recognises a mouse
ZnT8 epitope as well as a human one. The alternative ZnT8 antibody available in the
lab was not species specific so could not be of use either.
An anti-c-myc antibody was also tested using cytochemical analysis of isolated islets
to detect the epitope tag also included at the 3’ end of the ZnT8 transgene. However
the staining was very weak with no real positive cells and just a faint background
despite trying several concentrations of the antibody (results now shown).
122
5.4 Discussion
In this Chapter our aim was to overexpress ZnT8 in the pancreatic α-cell in order to
investigate the effect this would have on glucagon secretion. The results, which are
reminiscent of our findings in αZnT8 KO animals (Chapters 2,3), reveal that
overexpression of ZnT8 in the α-cell has no effect on glucose homeostasis or insulin
sensitivity (assessed by intraperitoneal tolerance tests), but does affect
hypoglycaemia responses as studied in clamps, and glucagon secretion in response
to low glucose both in vivo and in vitro. The findings from the transgenic mouse with
respect to glucagon release are thus reciprocal to those obtained from αZnT8 KO
mice and further reinforce the view that ZnT8 has an important role in the α-cell in
regulating glucagon secretion, at least in the hypoglycaemic state.
The efforts made here to quantify the degree of human ZnT8 expression in the α-cell
at the protein level were, however, unsuccessful. This is, of course, challenging given
that α-cells comprise only a small fraction of the rodent islet [318], requiring single
cell analysis either after FACS or the use of imaging approaches. Had there been
more time, other anti-human ZnT8 antibodies would have been tested in order to
determine the expression of human ZnT8 at the protein level and not just the at the
mRNA level. The fact that expression at the mRNA levels is relatively high compared
to the endogenous control gene lead to the expectation of high enough protein
levels that would be detected by immunohistochemistry. FACS would be even more
sensitive at detecting positive cell populations. It is thus perhaps just an issue of
finding the appropriate antibody.
The results here complement those of a similar study from the laboratory [317]. In
this earlier study, human ZnT8 was inducibly overexpressed in the adult β-cell by
using an insulin promoter-dependent Tet-On system. Glucose-induced insulin
secretion from the βZnT8Tg mice was impaired and Zn2+ release during stimulated
123
exocytosis was elevated. The same would be expected from the α-cell considering
how deletion of ZnT8 led to a reduction of granular Zn2+ levels in both the α-cell and
β-cell. Further investigations will be required to test this possibility.
In this earlier study, however, the effect of deletion and overexpression of ZnT8 on
insulin secretion were not reciprocal as we observed in the α-cells. Although various
groups [222, 252, 268] have shown an impairment in insulin secretion following ZnT8
ablation, no change was detected in the levels of insulin released. By contrast, Zn2+
release respectively increased and decreased in βZnT8Tg and βZnT8KO mice [317].
5.5 Conclusion
Overexpression of ZnT8 in α-cells, in common with its deletion, affects glucagon
release at hypoglycaemic conditions in this case causing a reduction in secretion.
ZnT8 in the α-cell is thus necessary for normal glucagon responses in the context of
hypoglycaemia. Both rare [221] and common variants [203] in the human SLC30A8
gene may thus lead to changes in both insulin and in glucagon release to affect
glucose homeostasis and T2D risk.
124
Chapter 6
Final Discussion
125
6.1 Investigating the role of ZnT8 in the pancreatic α-cell through mouse
molecular genetics
The aim of this thesis was to investigate the role of the Zn2+ transporter ZnT8 in the
α-cell and the regulation of glucagon secretion. The approach here was to generate
mice deleted for or overexpressing ZnT8 specifically in the α-cell. Although ZnT8 has
been widely studied in the β-cell [271, 272] this is the first time that it has been
extensively investigated in the α-cell. The β-cell findings from the various groups
researching ZnT8 have been controversial but strongly imply a role for ZnT8 in β-cell
Zn2+ homeostasis, insulin packaging and insulin secretion. It was thus important to
understand whether ZnT8 is important for α-cell function. A SNP in the SLC30A8
locus encoding ZnT8, resulting in a transporter with reduced transporting ability, was
found to be associated with increased T2D risk [203]. More recently, rare loss-of-
function ZnT8 variants [221] were found to be protective against T2D in contrast to
the previous findings [203]. Understanding the function of ZnT8 not only in the β-cell
but also in the α-cell where it is highly expressed may thus provide some insight and
help determine how and why specific ZnT8 variants affect T2D risk.
T2D is a polygenic disease with many genes possibly contributing to disease
susceptibility and pathogenesis. Gaining knowledge about the role of T2D associated
genes in pancreatic islets could lead to the identification of potential targets for the
development of anti-diabetic drugs.
The ability to generate cell type-specific and whole body knockout mice has made
these an instrumental component in studies investigating the role of a protein in the
whole body or in a certain cell type. Results from rodent studies can be used to
deduce the likely effects a disease-associated gene might have on human physiology.
This method was utilized in this project in an effort to determine whether changes in
126
the activity of ZnT8 in the α-cell could contribute to the effects of ZnT8 risk alleles in
humans.
6.2 ZnT8 in the pancreatic α-cell
In agreement with earlier findings [222] we showed that although ZnT8 is highly
expressed in the α-cell, colocalization with glucagon is incomplete even more so than
in the β-cell. Thus, ZnT8 appears to reside in cellular locations other than glucagon
granules (Figure 3.1). Although the identity of such structures has not been explored
here a possibility is that they may at least partly be immature granules. Future
studies using higher resolution techniques (including super-resolution techniques
such as PALM/STORM) or immuno-electron microscopy will be needed to provide a
definitive answer.
To address the question regarding the role of ZnT8 in the α-cell and glucagon
secretion we began by generating a mouse that lacked ZnT8 specifically in the α-cell
only. Reports from other studies using the same PPG.Cre line that was used here
showed a range of recombination levels from 13% [298] to 45% [300] so it was
important to determine the extent of deletion in our mouse. FACS analysis revealed
that the recombination rate was quite low; with an overall ~15% reduction in ZnT8 in
the entire α-cell population. However, no deletion was observed in the β-cell
population demonstrating the selectivity of this approach.
Thus the observed effects may have been dampened compared to having had
complete or near complete ablation of ZnT8. Additionally, this could mean that any
subtle changes occurring due to the absence of ZnT8 might not have been detected
due to the low level of deletion achieved.
127
αZnT8 KO mice displayed normal glucose tolerance as assessed by glucose tolerance
tests, and normal insulin sensitivity. There were no changes in fasting blood glucose
or fasting plasma glucagon levels. However, there was an effect on the release of
glucagon under low glucose conditions. Deletion of ZnT8 caused an increase in
glucagon secretion at hypoglycaemic conditions both in vitro and in vivo. These
changes in glucagon release cannot be accounted for by increased glucagon content
in KO mice since total pancreatic glucagon measured by an immunoassay was
unchanged as were glucagon mRNA levels. An increase in α-cell area of ~15% was
observed in the KO pancreas and this could be contributing to the enhanced
glucagon release but the increase was too small to account for the 2 fold change
detected in glucagon secretion by KO islets. No changes were observed in Ca2+
dynamics as peak amplitude and frequency of oscillations were comparable between
the two cell populations, WT and KO. Thus, the mechanism by which ZnT8 influences
glucagon release is Ca2+ independent.
We showed a reduction in granular Zn2+ content in KO cells suggesting that ZnT8 is
involved in the accumulation of Zn2+ in glucagon granules. Based on the decreased
Zn2+ content [268] and the findings of a loss of Zn2+ co-release with insulin when
ZnT8 is deleted in the β-cell [270], an elimination of Zn2+ from granules upon
exocytosis is to be expected from α-cells following ZnT8 ablation. The impaired
release of Zn2+ from α-cells could be involved in a mechanism via which ZnT8
suppresses glucagon release at low glucose concentrations. The likely reduction in
local Zn2+ levels could also be affecting secretion.
There have been several rodent studies looking into the effect of Zn2+ on glucagon
release, the vast majority demonstrating an inhibitory action of Zn2+. The Zn2+
chelator Ca2+-EDTA caused stimulation of secretion from the perfused rat pancreas
[84]. Incubation of isolated rat cells with Zn2+ diminished pyruvate and glucose
induced glucagon secretion through the reversible activation of KATP channels and
128
subsequent reduction in electrical activity [121]. A similar study in the mouse
showed an inhibitory effect of Zn2+ on glucagon release on an α-cell line and isolated
islets with maximal inhibition at 10 µM Zn2+ [130]. This inhibitory action on glucagon
secretion was confirmed by another group experimenting with whole islets and FACS
sorted α-cell at low glucose concentrations [320]. However, the above effect of Zn2+
in the mouse cannot be attributed to KATP channel activation. Instead, it involved the
uptake of the ion by Ca2+ channels and a change in the redox state of the cell [130].
Thus the likely reduction in Zn2+ released from secretory granules could be alleviating
the suppression normally imposed by the ion on surrounding α-cells and could be
contributing to the enhanced glucagon secretion observed from KO islets (Figure
6.1).
Correspondingly, overexpression of ZnT8 led to a reciprocal decrease in glucagon
secretion from isolated islets at low glucose (Chapter 5). This further confirms the
importance of ZnT8 in maintaining normal α-cell responses during hypoglycaemia.
Research on mouse cell lines are in agreement with the findings in this study.
Overexpression of ZnT8 variants resulted in a reduction in glucagon secretion by 50%
while knockdown caused an increase in secretion by 70% [274]. The changes in
glucagon secretion were accompanied by corresponding changes in glucagon
content which was not observed in our study since total glucagon content was
unchanged.
Zn2+ is important in the β-cell for insulin crystallization and storage in the granules
[267]. No such role for Zn2+ in the α-cell has been described yet the presence of Zn2+
in glucagon granules has been confirmed by electron microscopy although the
staining is less intense that in insulin granules [302]. It is thus plausible that Zn2+ may
be involved in glucagon packaging within the granule. A less compact aggregation of
individual glucagon particles inside the granule could enable the release of the
peptide via the dilating fusion pore so that a bigger fraction of exocytotic events go
through to completion prior to pore closure [321].
129
A
B
C
Figure 6.1 – Hypothetical model reconciling data on the effect of Zn2+ on glucagon
secretion and our mouse models. (A) Glucagon secretion occurring at normal levels from
WT α-cells with Zn2+ released from the granules during exocytosis. Zn2+ can enter
surrounding cells where it can influence glucagon secretion in an inhibitory manner. (B) α-
cells lacking ZnT8 have lower levels of Zn2+ in their secretory granules and subsequently
lesser amounts are released in the extracellular space. Less Zn2+ enters neighboring cells so
inhibition of glucagon secretion is lower thus more glucagon is released. (C) In ZnT8
overexpressing α-cells, secretory granules are expected to contain more Zn2+ meaning more
would be released in the extracellular space and more would enter the surrounding cells
imposing more inhibition on glucagon release. That would explain the lower levels of
glucagon secretion observed.
130
The full effect of a complete loss of ZnT8 may not be apparent due to compensatory
gene expression changes possibly during development which have not yet been
identified. The obvious candidates are the other members of the ZnT family of
transporters. However, arguing against this mechanism gene expression analysis
showed the presence of mRNAs encoding several ZnTs in the mouse α-cell but there
was no evidence for a compensatory change in their expression levels in islets with
α-cell specific deletion of ZnT8. Similarly there was no compensatory increase in
expression following global [222] or β-cell specific ablation of ZnT8 [270, 272]. On
the other hand findings from knockdown studies in cell lines and KO mouse models
indicate that ZnT3 [322, 323] and ZnT7 [324] could be involved in the control of
glucose stimulated insulin secretion in a ZnT8 independent manner at least in mice.
ZnT7 deletion caused decreased insulin secretion and increased insulin resistance
leading to glucose intolerance [324]. Of note, however multiple studies form our
laboratory have failed to measure detectable levels of ZnT3 mRNA in mouse islets
[222, 325].
The observed enhancement in glucagon secretion was achieved when ZnT8 was
deleted from only a small proportion of α-cells. This indicates that the absence of
ZnT8 has a large effect at the level of individual α-cells or that the KO cells may have
a disproportionate role in regulating the activity of the rest of the α-cell population.
The data gathered from this study does not eliminate either possibility. It is unlikely
that this increase in total secretion is achieved by an elevation of release at the level
of individual α-cells lacking ZnT8 since they would have to secrete ~7 fold more to
achieve a 2-fold change in glucagon levels.
Another possibility is that α-cells are highly inter-connected (and therefore inter-
dependent) as reported by our laboratory to be the case for β-cells [326]. The islet
with its 3D structure is thought to be conducive to intercellular signaling for
coordination of insulin secretion. The distinct flow of information through
131
interconnected cells causes an organized release of insulin responding to glucose or
other mediators.
6.3 ZnT8 and Zn2+ homeostasis in the α-cell
Cytosolic free Zn2+ measurements were performed using the molecular Zn2+ probe
FluoZin-3 and revealed significant reductions in cytosolic levels following ZnT8
ablation (Chapter 4). Considering that ZnT8 was only deleted in 50% of the RFP
population it is possible that the lowering of Zn2+ levels was underestimated. A
decrease in Zn2+ was also detected in the granules using ZinPyr-4 a molecular probe
that localizes to the granules. These effects of ZnT8 deletion are consistent with
observations in the β-cell [222, 268]. The reduction in granular Zn2+ levels is in
agreement with the proposed role of ZnT8 as an efflux transporter on granule
membranes. On the other hand, the decrease in cytosolic Zn2+ is less easy to explain
but may reflect a decrease in cytosolic Zn2+ uptake across the plasma membrane.
Consideration of the biology of another member of the ZnT family, ZnT5, suggests an
alternative possibility. ZnT5 is ubiquitously expressed and more abundant in islets
compared to other tissues. It has been suggested that it may have the ability for
bidirectional transport [311]. It seems to have discrete roles in Zn2+ homeostasis
since in addition to being expressed on secretory granules and the Golgi apparatus in
the β-cell, [247] it was found to be expressed on the plasma membrane where it is
thought to mediate Zn2+ influx instead of efflux [311]. ZnT8 immunostaining was
detected at the plasma membrane as well as secretory granules [222]. A similar role
in influx as ZnT5 at that location could explain the reduction in cytosolic Zn2+ levels
observed both in α-cells and β-cells following cell specific deletion of ZnT8.
Transfection of HeLa cells, which do not normally express ZnT8, with a ZnT8
construct led to an increase in cytosolic Zn2+ [213]. This finding is also consistent with
a role for ZnT8 in Zn2+ influx.
132
6.4 ZnT8, SLC30A8 and T2D risk
The mechanism involved in driving the association between rs13266634 risk alleles
and T2D has previously been investigated. Nicolson et al [222] observed the
accumulation of Zn2+ in the cell using molecular probes following transient
expression of either the Arg or Trp variants in a β-cell line. Cells expressing the Trp
construct had a greater fluorescence change suggesting that this variant is a more
active transporter. How this change, i.e. increased cytosolic free Zn2+ is actually
beneficial to the β-cell is still unclear since the Trp variant is not linked to improved
basal or glucose stimulated insulin secretion from human islets [327]. In addition the
report by Flannick et al [221] that haploinsufficiency for SLC30A8 is associated with
significantly (>50%) reduced risk of T2D contradicts the finding of a slightly less
active ZnT8 causing an increase in T2D risk. Thus it appears that a more intricate
mechanism is accountable for the rs13266634 association to T2D and the role of
ZnT8 in the control of hormone release [328].
The complexity regarding the role of ZnT8 could at least partly involve the
interaction between the pancreas and the liver. Zn2+ co-secreted with insulin plays a
part in the regulation of hepatic hormone clearance [271] where is suppresses
uptake by inhibiting clathrin-dependent endocytosis.
Taking into consideration the effect of haploinsufficiency potential explanations are
that the Trp variant turns over faster than the Arg one or that it traffics to different
sub-cellular locations [223]. The latter seems likely since the biggest difference
between the Arg and Trp variants was observed during measurements of
cytoplasmic Zn2+ accumulation [222] suggesting ZnT8 can transport Zn2+
bidirectionally being involved in influx across the plasma membrane. A possibility is
that the protective effect of the Trp variant may arise from increased activity at the
plasma membrane although how this is protective is as yet unclear [223].
133
A hypothetical model has been proposed by Davidson and colleagues [223] to bring
together data for rare and common SLC30A8 variants. Glucose stimulated insulin
secretion leads to an elevated production of reactive oxygen and nitrogen species
[329]. Taking into account the fact that Zn2+ plays a key part in moderating oxidative
stress [330] a theory that could connect the contrasting observations is that the Trp
and loss-of-functions variants work by briefly delivering more Zn2+ into the
cytoplasm during secretion (Figure 6.2). This increase in Zn2+ in turn might reduce
the deleterious effects of reactive oxygen species. The more active Trp variant at the
plasma membrane would increase cytosolic Zn2+ by an increased reuptake of Zn2+ co-
secreted with insulin from the extracellular space. In the case of the loss-of-function
variants, their presence at the granule membrane would similarly lead to an increase
in cytosolic Zn2+ through a reduction in transport of the ion in the granule lumen
[223]. This is a model suggested for the β-cells but something similar could be
occurring in the α-cell to explain the observations on T2D risk.
Although plausible there is a weakness in this scheme since it implies that a loss of
function variant would actually lead to an increase in cytosolic Zn2+ levels whereas in
KO studies the opposite has been observed. In the α-cell we showed a reduction in
cytosolic Zn2+ following ZnT8 ablation. Similarly β-cell specific deletion led to a
reduction in intracellular Zn2+ levels [222, 268]. Thus it appears that the relationship
between the ZnT8 variants and T2D risk is more complicated.
134
Figure 6.2 – Hypothetical model reconciling data for rare and common SLC30A8 variants in
the β-cell. The speculation is that the rs13266634 allele encoding the 325 Trp variant and
the rare mutations leading to haploinsufficiency are protective due to prolonging the
transient elevation in cytosolic Zn2+ levels occurring during glucose stimulated insulin
secretion. This subsequently reduces the harmful effects of oxygen reactive species. In (A)
the 325 Trp variant increases cytosolic Zn2+ levels by enhanced reuptake of Zn2+ secreted
alongside insulin at the plasma membrane. In (B) rare loss of function mutations lead to an
increase in Zn2+ levels primarily by reduced uptake of Zn2+ into secretory granules. Other ZnT
family members could be compensating for the possible decrease in ZnT8 facilitated uptake
at the cell membrane. Adapted from Davidson et al [223].
A
B
135
Considering that the activity of the Arg form of ZnT8 is 30% less than the Trp variant
[259], it is reasonable to assume that Zn2+ accumulation into granules would be
lowered by 15% and 30% in heterozygote and homozygote Arg allele carriers
respectively. Thus heterozygote loss-of-function would have a 50% reduction [328].
Data from mouse studies primarily suggest a harmful effect on glucose homeostasis
when ZnT8 is deleted in the β-cell yet it is likely that there are species-specific
differences in the role of ZnT8 and Zn2+ in the islet and in hepatic hormone clearance
[271]. It is likely that in humans the interplay between ZnT8 activity and disease risk
may involve a complex dose-response. Interplay between actions on α- and β-cells
may contribute to this complexity.
In this case even small reductions in ZnT8 activity, for example the Arg325 variant,
could be deleterious while in the case of the loss-of-function variant a more sizeable
reduction could be protective in a situation where enhanced insulin secretion offsets
the harmful effect of elevated clearance [328]. Conversely in the mouse there may
be a less complex relationship where elevated hepatic clearance as a result of falling
ZnT8 levels leads to increasingly impaired insulin action.
More recent studies seek to determine not just which loci are associated with T2D
but how such loci are associated with glycaemia and disease pathogenesis. Co-
expression analysis demonstrated that the expression of many genes correlated
strongly with glucagon. One of these genes was SLC30A8. The lower the ZnT8
expression was in human islets, the lower the expression of glucagon [331]. Such a
finding suggests that ZnT8 may be involved in the control of glucagon expression or
vice versa. This is not something we however detected in the mouse. Although the
KO islets expressed less ZnT8 the expression of glucagon did not differ significantly
when compared to WT islets.
136
In addition, SLC30A8 was found to be a target for the transcription factor 7-like 2
(TCF7L2) by our laboratory [332], a finding later confirmed by others [331]. Variants
of this transcription factor are associated with increased risk of T2D, having the
highest effect on disease risk out of all the T2D variants uncovered so far by GWAS
[207]. It is thus possible that ZnT8 is involved in a regulatory network with other
diabetes-associated genes [333] so that specific variants of such genes may regulate
α- and β-cells and thus have an effect on insulin and glucagon levels and
consequently T2D risk.
6.5 Final conclusions
By creating tissue specific KO mice where ZnT8 is deleted specifically in α-cells, and
transgenic animals overexpressing this transporter inducibly in these cells, we
demonstrate that the Zn2+ transporter regulates normal glucagon secretion during
hypoglycaemia. Additionally we show that ZnT8 is important for intracellular Zn2+
homeostasis affecting both cytosolic and granular free Zn2+ levels. Changes in ZnT8
expression or activity in the α-cell may thus contribute to diabetes risk in SLC30A8
risk carriers. ZnT8 might therefore provide a suitable target for pharmacotherapeutic
interventions acting to modulate both insulin and glucagon secretion in the context
of both T2D and glucagon secretion during hypoglycaemia in T1D.
137
6.6 Future directions
Future studies are needed to provide further insight into the role of ZnT8 in the α-
cell and to complement the findings from this study.
One of the major effects ZnT8 had on the β-cell was on the morphology of secretory
granules. It would be possible to investigate this in the α-cell by staining for glucagon
in order to identify them during electron microscopy.
RNA seq analysis of RFP+ cells from WT and KO islets would give information into
whether ZnT8 deletion causes any specific changes to the transcriptome in the α-
cells and whether any such changes could account for the observed differences in
glucagon secretion.
To assess the theory of Zn2+ contributing to the observed effects of deletion of ZnT8
on glucagon secretion we could use the Zn2+ chelator TPEN to check if lowering Zn2+
levels in the surrounding of the cells could further enhance the exaggerated
glucagon response in low glucose concentrations. Alternatively we could use the Zn2+
ionophore pyrithione to evaluate whether increased entry of the ion into the cell
dampens the elevated glucagon levels observed during hypoglycaemia in ZnT8 KO
mice.
Crossing the ZnT8TgGlurtTA mice with a mouse containing a fluorescent reporter
under Tet On control would allow its expression specifically in the α-cell allowing
experiments at the single cell level as those performed in the ZnT8 KO mice. It would
be interesting to see whether the ZnT8 overexpressing α-cells have a reciprocal
change in intracellular Zn2+ levels.
138
Chapter 7
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Publications
Gerber, P.A., Bellomo, E.A., Meur, G., Hodson, D.J., Solomou, A., Hollinshead, M., Chimienti, F., Lavallard, V.J., Bosco, D., Hughes, S.J., Johnson, P.R. and Rutter, G.A. Hypoxia alters the expression of the type 2 diabetes-associated Zn2+ transporter Slc30a8/ZnT8 and cytosolic Zn2+ levels in human and rodent islets Diabetologia, 57(8):1635-44 (2014)
Sun, G., da Silva Xavier, G., Gorman, T., Priest, C., Solomou, A., Hodson, D.J., Foretz, M., Viollet, BN., Herrera, P-L., Parker, H., Reimann, F., Gribble, F.M., Migrenne, S., Magnan, C., Marley, A. and Rutter G.A. LKB1 and AMPKa1 are required in pancreatic alpha cells for the normal regulation of glucagon secretion and responses to hypoglycemia. Mol Metab 4(4):277-286 (2015)
Solomou, A., Meur, G., Bellomo, E., Hodson, D.J., Tomas, A., Migrenne Li, S., Philippe, E., Herrera, P., Magnan, C., and Rutter G.A. The zinc transporter Slc30a8/ZnT8 is required in a subpopulation of pancreatic α cells for hypoglycemia-induced glucagon secretion. J Biol Chem 290(35):21432-21442 (2015)
Akalestou, E., Christakis, I, Solomou, A.M., Minnion, J.S., Rutter, G.A. and Bloom, S.R. Proglucagon-derived peptides do not significantly affect acute exocrine pancreas secretion in vitro and in vivo in the rat. Pancreas, in press (2015) Rutter, G.A., Chabosseau, P., Bellomo, E.A., Maret, W., Mitchell, R.K, Hodson, D.J., Solomou, A., and Hu, M. Intracellular zinc in insulin secretion and action: a determinant of diabetes risk? Proc Nutrition Soc. in press (2015)
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