The Role of SIRT1 in Pancreatic Beta Cells

by

Lemieux Luu

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Physiology University of Toronto

© Copyright by Lemieux Luu 2013

ii

The Role of SIRT1 in Pancreatic Beta Cells

Lemieux Luu

Master of Science

Department of Physiology University of Toronto

2013

Abstract

SIRT1 has emerged as a critical regulator of glucose homeostasis and metabolism in the past

decade. Glucose homeostasis is tightly regulated by insulin however, the factors affecting

insulin release are still incompletely understood. Relatively recent evidence has shown SIRT1 to

be a positive mediator of insulin secretion although its mechanism is largely unknown.

Therefore, the aim of this study was to determine how SIRT1 regulates insulin release. Using a

pancreatic beta cell-specific Sirt1 knockout mouse model (Sirt1BKO), oral glucose challenge

revealed a glucose intolerant phenotype with reduced insulin secretion. Isolated Sirt1BKO islets

also secreted less insulin without changes to insulin content or islet morphology. Intracellular

defects were localized to the mitochondria and showed suppressed bioenergetics negatively

affecting downstream glucose-induced calcium influx. This is the first study using a Sirt1BKO

mouse model to show novel mitochondrial genes under SIRT1 regulation and when impaired,

results in reduced insulin secretion.

iii

Acknowledgments

I would like to express my sincere gratitude to my supervisor, Dr. Michael B. Wheeler, for taking

me into his productive lab and giving me the opportunity to work on this exciting project. He

has provided me with endless guidance throughout and his mentorship has brought my

scientific calibre to a new level. His training has armed me with the confidence to tackle the

unknowns of science with poise and I am thankful for all of my experiences in his lab.

I would also like to acknowledge my lab colleagues (past and present): Dr. Alexandre Hardy, Dr.

Alpana Bhattacharjee, Chinyee Huang, Dr. Christina Basford, Dr. Christine Robson-Doucette, Dr.

Emma Allister, Dr. Fay Dai, Dr. Jakob Bondo Hansen, Dr. Junfeng Han, Kacey Prentice, Dr. Ming

Zhang, Sandro Serino, and Dr. Ying Liu. The long hours and arduous process of research

becomes a little more bearable with you all by my side. You have all provided me with

tremendous support, knowledge, and importantly, laughter and I’ll always remember you all.

I want to thank my committee members: Dr. Adria Giacca and Dr. George Fantus for their

invaluable insight into my project which has steered me towards success. I would like to thank

our collaborator, Dr. Jamie Joseph for welcoming into his lab to perform the lovely Seahorse

experiments that aided critically in my manuscript.

To my girlfriend, Dianna, thank you for your unending patience and love. For all the evenings

and weekends that the lab stole me away from you, you understood and rooted for me every

step of the way. I’m lucky to have you in my life. And lastly but most importantly, I want to

thank my family. Your unconditional, unwavering love and support has made all of my

achievements possible. Your complete and utter belief and faith in me has given me the

iv

courage to tackle all of the hardships, obstacles and adversity I’ve faced in life. To my brother

and sister, Alex and Diana, you two have played such an enormous part in molding me into the

person I am today on top of everything else you both do for me. I couldn’t ask for better

siblings. And to my mother and father, Hanh and Le, your boundless encouragement and love is

immeasurable and my appreciation for you both is indescribable. You’ve taught me things that I

can’t learn in a lab or library, taught me the important things in life. You’ve both given me

everything and I can only hope to repay you by making you proud each and every day.

v

Publications that Contributed to this Thesis

Luu, L., et al., The loss of Sirt1 in mouse pancreatic beta cells impairs insulin secretion by disrupting glucose sensing. Diabetologia. 2013 Apr 29. PMID: 23783352

vi

Table of Contents

Abstract………………………………………………………………………………………………………..….……………………ii

Acknowledgements……………………………………………………………………………………………………..……….iii

Table of contents………………………………………………………………………..…………………………………....….vi

List of Figures……………………………………………………………………………………………………………….…….viii

List of Abbreviations……………………………………………….…………………….……………………………………..ix

1 Introduction

1.1 Type II Diabetes and the Pancreatic Beta Cell…………………………..….……..………....…1

1.1.1 Type 2 Diabetes……………………………………………………………………………….….….1

1.1.2 Pancreatic Beta Cells and Glucose-stimulated Insulin

Secretion…………………………………………………..………………..….………………….…..2

1.1.3 Mitochondrial and Insulin Secretion…………………………………………….………...6

1.2 Introduction to SIRT1………………………………………………………………………………….……11

1.2.1 Sirtuin Family………………………..…………..…………………………………………….…...11

1.2.2 SIRT1 Biological Function and Targets…………..………………………………………13

1.2.3 SIRT1 and the Pancreatic Beta Cell …..…………………………….…………………….19

1.2.4 Sirtuins and Insulin Secretion ………………………………………………..……..………22

2 Rationale and Hypothesis………….…………………………………………………………………………….….24

3 Materials and Methods……………………………………………….………………………………………………26

3.1 Experimental SIRT1 Inactivation Models……………………………………………………………..26

3.1.1 Sirt1BKO Mice…………………………………………………………………………………..….26

3.1.2 SIRT1KD MIN6 Cells………………………….…………………………………………………..27

vii

3.2 Glucose and Insulin Tolerance Tests…………………………………….…………….…..…………..27

3.3 Islet Isolation and Dispersion………………………………………….………………….…..….……….28

3.4 Glucose- and KIC-stimulated Insulin Secretion and htrf.…………..……………….…………28

3.5 Western Blot……………………………………………………………………………………………………….29

3.6 Polymerase Chain Reaction and Microarray…………………….……………………….…………29

3.7 Immunohistochemistry……………………………………………………………………………….……...30

3.8 Transmission Electron Microscopy………………………………………………………….…….…….31

3.9 Calcium Imaging and Mitochondrial Membrane Potential……………………..……………31

3.10 Oxygen Consumption Rate………………………………………………………….………………..…….32

3.11 ATP Measurements……………………………………………………………….……….…………………..32

4 Results………………………………………………………………………………………………………………..………33

4.1 Validation of SirT1 Inactivation……………………………………………………………………………33

4.2 SirT1BKO Mice are Glucose Intolerant……………………………………………………….….…….35

4.3 SIRT1 Inactivation Reduces Glucose-stimulated Insulin Secretion……..………………..36

4.4 The Effects on Insulin Secretion are Due to Sirt1 Inactivation ……………………….…...38

4.5 Defects in Insulin Secretion are Downstream of Glycolysis……………..………….………39

4.6 Sirt1 Knockdown Causes Significant Dysregulation of Metabolic Genes……….…….40

4.7 SIRT1 Inactivation Impairs Mitochondrial Function……………………………………………..43

4.8 ATP Production is Impaired by Pharmacological SIRT1 Inhibition……………..…………45

4.9 Decreased Calcium Influx in Sirt1BKO Cells………………………………………….……………..45

5 Discussion…………………………………………………………………………………………………………….…….48

6 Conclusions……………………………………………………………………………………………………….………..57

7 References………………………………………………………………………………………………………………….58

viii

List of Figures

Figure 1: Schematic representation of beta cell glucose-stimulated insulin secretion…….…3

Figure 2: The electron transport chain located in the inner membrane…………………………….5

Figure 3: Role of mitochondria and insulin resistance……………………………………………………….8

Figure 4: Schematic representation of SIRT1-mediated substrate deacetylation……………..12

Figure 5: Schematic representation of selected SIRT1 targets…………………………………….…..16

Figure 6: TNFa ligand binding to the TNF Receptor leads to activation of JNK1………..……..18

Figure 7: Schematic of tamoxifen-inducible CreER Lox system…………………………………………26

Figure 8: The inactivation of Sirt1 in Sirt1BKO mice……………………………………………………..….34

Figure 9: Impaired glucose tolerance and decreased insulin secretion in Sirt1BKO mice...35

Figure 10: Decreased insulin secretion in Sirt1BKO islets…………………………………………………..37

Figure 11: SIRT1 is responsible for changes in insulin secretion………………………………..……….38

Figure 12: Reduced alpha-ketoisocaprioic acid-induced insulin secretion in Sirt1BKO mice………………………………………………………………………………………………………………....40

Figure 13: Dysregulation of mitochondria-related genes in SIRT1KD MIN6 cells………….…….41

Figure 14: Sirt1 inactivation does not affect mitochondrial mass……………………………..……..42

Figure 15: Sirt1 inactivation impairs mitochondrial hyperpolarization and OCRs in islets from Sirt1BKO and SIRT1KD mice. ……………………………………………………………………44

Figure 16: Decreased ATP production and calcium influx in Sirt1-inactivated cells…….……..46

ix

List of Abbreviations

BESTO Beta Cell-specific Overexpressing

CreER Inducible Cre Recombinase

DNP 2,4-Dinitrophenol

ETC Electron Transport Chain

GLP-1 Glucagon-like Peptide-1

GSIS Glucose-stimulated Insulin Secretion

HTRF Homogeneous Time Resolved Fluoresence

ITT Insulin Tolerance Test

KIC Alpha-ketoisocaproate

KRB Krebs-Ringers Bicarbonate Buffer

MIN6 Clonal Mouse Insulinoma Pancreatic Beta Cell

MMP Mitochondrial Membrane Potential

NADH/NAD+

NAFLD Non-alcoholic Fatty Liver Disease

Nicotinamide Adenine Dinucleotide

NAM Nicotinamide

OCR Oxygen Consumption Rate

OGTT Oral Glucose Tolerance Test

OXPHOS Oxidative Phosphorylation

RIA Radioimmune Assay

ROS Reactive Oxygen Species

SirBACO SirT1 Bacterial Artificial Chromosome Overexpressor

Sirt1BKO Pancreatic Beta Cell-specific Sirt1 Knockout

SIRT1KD Sirt1 Knockdown

STZ Streptozotocin

TCA Tricarboxylic Acid

1

1 Introduction

1.1 Type II Diabetes and the Pancreatic Beta Cell

1.1.1 Type II Diabetes

It is estimated that nearly 350 million individuals worldwide are afflicted with

diabetes[1] and its complications are the leading cause of death around the globe

(http://who.int/mediacentre/factsheets/fs310/en/). Diabetes comprises a group of metabolic

disorders which include chronically high blood glucose, defective insulin secretion and/or

action[2]. While diabetes falls into two broad categories: type 1 (near total loss of insulin-

producing cells) and type 2 (dysfunctional beta cells), the main focus here will be type 2

diabetes (T2D) which accounts for more than 90% of all diabetic cases[3]. The causes of T2D are

multi-factorial and complex although what is understood is that diabetes begins with

hyperglycemia and abnormal fasting plasma glucose[4-6]. The American Diabetes Association

defines this stage as “pre-diabetes”[6] and it is highly associated with obesity and a sedentary

lifestyle[7]. Without altering one’s unhealthy habits, studies show the progression towards

diabetes can occur in as few as 29 months[6]. This process is characterized by somewhat

discrete stages beginning with hyperinsulinemic compensation from beta cells to counter the

hyperglycemia due to insulin resistance[8, 9]. This stage can persist for years until the acute

insulin response of beta cells becomes impaired[10] and lays the way for the next steps towards

frank diabetes. Although the exact mechanism for the progressive decline in beta cell function

2

isn’t clear, what’s known is these events correlate precisely with abnormally high glucose

levels[11] as well as excessive fatty acids[12]. This can be combined with decreased beta cell

mass exacerbating the pathology[13]. Eventually individuals with diabetes can no longer

secrete adequate amounts of insulin to regulate their blood levels, leading to increased levels

of morbidity and mortality.

It may come as a surprise that while the beta cell’s ability to secrete insulin is central to overall

glucose homeostasis and the dysfunction of this cell shifts the balance towards development of

type 2 diabetes, our understanding of the underlying mechanics of glucose-stimulated insulin

secretion (GSIS) is incomplete. Therefore, the following sections will elaborate on the findings

and unknowns of GSIS.

1.1.2 Pancreatic Beta Cells and Glucose-stimulated Insulin Secretion

Understanding the beta cell is imperative to understanding the etiology of diabetes. This cell

type plays a unique role in the body as it serves not only to augment rises in blood glucose

levels, but it also helps to maintain basal glucose levels[14]. At rest, a human being’s glucose

concentration is approximately 5mmol/l[14]. Upon ingestion of a meal, glucose levels rise in the

blood and cause increased glucose uptake into the beta cell via the GLUT-2 in mice or the GLUT-

1 and GLUT-3 transporters in humans[15]. Once inside the cell, glucose encounters Glucokinase

(GK) which acts as the predominant beta cell glucose sensor, a protein which allows for

immediate and accurate quantification of glucose levels[16]. GK phosphorylates glucose to

form glucose-6-phosphate (G6P) and this molecule can then undergo metabolism by entering

3

glycolysis[17]. Glycolysis is a series of 10 enzyme-catalyzed reactions which converts G6P into

two molecules of H20, two molecules of ATP (net), two molecules of NADH, and two molecules

of pyruvate[18]. The pyruvate formed enters the mitochondrial matrix where it is converted

into Acetyl-CoA, CO2, and once again, NADH[19]. While the details of the TCA cycle are beyond

the scope of this thesis, a noteworthy product of this process is the production of three NADH

molecules. The accumulated NADH molecules proceed to act as reducing agents in a process

called oxidative phosphorylation.

Figure 1: Schematic representation of beta cell glucose-stimulated insulin secretion. i: glucose uptake into the beta cell. ii: formation of proton gradient via mitochondrial electron transport. iii: Oxidative phosphorylation and rise in ATP/ADP. iv: Katp channel closure and subsequent membrane depolarization. v: activation of voltage-dependent calcium channels (VDCC) leading to calcium influx. vi: calcium-triggered insulin secretion.

4

Oxidative phosphorylation (OXPHOS) is a metabolic process that occurs in the

mitochondria whereby organisms harvest energy from nutrients to form ATP[20]. The process

comprises a series of mitochondrial intermembrane-embedded protein complexes which act as

protein acceptor/donors in redox reactions[21]. The release of energy from the redox reactions

or so-called electron transport chain (ETC) creates potential energy in the form of a proton

gradient[22, 23]. As a result of this highly efficient process, oxidative phosphorylation is largely

conserved from prokaryotes to eukaryotes[24, 25].

To begin, the NADH formed from glycolysis and the TCA cycle donates a pair of electrons to the

first protein complex in the chain, NADH Ubiquinone Oxidoreductase (Complex I)[26].

Mammalian Complex I is a large protein (~100kDa) consisting of 45 subunits which are encoded

both by the nuclear and mitochondrial genomes which are responsible for its assembly,

regulatory function and enzymatic activity[20, 27]. This entire protein assembly uses the energy

of redox cycling to pump protons out into the intermembrane space of the mitochondria[28].

The next member of the ETC is Complex III, Ubiquinone-cytochrome c Oxidoreductase,

also known as cytochrome bc1 (Complex II is involved in the TCA cycle). It is also a protein

complex consisting of 11 subunits including an evolutionary core responsible for its redox

reactions which further pumps proton into the intermembrane space [29], [30]. These electrons

are then transferred onto the next component, Complex IV, Cytochrome c Oxidase which also

performs redox reaction[31]. It translocates protons from the mitochondrial matrix into the

intermembrane space contributing to the proton motive force[32]. Of note, complex IV is the

final reducing agent in the chain whereby it donates its electrons to molecular oxygen, O2,

5

which combines with four H+’s to form two molecules of water[33, 34]. This property of oxygen

consumption can be used as a measure of mitochondrial respiration.

Figure 2: The electron transport chain located in the inner membrane. NADH is oxidized by Complex i to NAD+. Electrons from Complex i are donated to Complex ii. Complex ii is oxidized by Complex iii which in turn reduces Complex iv. Complex iv transports electrons to molecular oxygen to form H2

O. At each complex, the energy from redox reactions is used to pump protons into the intermembrane space thereby setting up a proton motive force (PMF). PMF is used by Complex v., ATP Synthase to drive the production of ATP from ADP + inorganic phosphate (Pi). Figure adapted from Lesnefsky, EJ., and Hoppel, CL. Ageing Res Rev. 5, 402-433 (2006).

The accumulation of protons will then be used by Complex V, the F1F0 ATP Synthase.

The ATP Synthase is composed of multiple subunits encoded by both the mitochondrial and

nuclear genomes with a molecular weight of more than 550kDa[20, 35]. At this juncture it is

important to note that the multi-subunit nature of these complexes gives rise to the possibility

6

that respiration can be impaired if any of these subunits are compromised either in function

and/or expression[20]. ATP Synthase acts as a molecular turbine spinning the potential energy

of the proton gradient to drive the production of ATP from ADP + inorganic phosphate (Pi)[36,

37]. This ATP energy can then be used by the cell to perform useful work. However, in the

pancreatic beta cell, ATP plays an additionally important function by closing the KATP

channels[38-40]. At rest, the beta cell is polarized to approximately -70mV but increasing

glucose to stimulatory levels will reduce the flow of K+ out of the cell[41]. This leads to

membrane depolarization which activates voltage-dependent calcium channels allowing for

influx of Ca2+

ions[42, 43]. Increased intracellular calcium triggers insulin granule fusion with the

plasma membrane and subsequent release of insulin into the blood stream[44, 45]. From this

scheme of the beta cell’s insulin secretory pathway, it becomes clear that many steps can affect

the efficiency of insulin secretion, especially those steps that go awry in disease. Notably

though, is the mitochondria as it is intimately involved in glucose metabolism.

1.1.3 Mitochondria and Insulin Secretion

The observation that T2D is strongly associated with inactivity gives rise to an important

link with the metabolism of nutrient sources and metabolic disorders. Perturbations in the

mitochondria’s ability to process fuels can affect whole-body glucose homeostasis and can

subsequently contribute to the development of T2D[46, 47]. Thus, the role of the mitochondria

in the pathogenesis of T2D has recently become an important and highly scrutinized topic.

7

To begin, the mitochondria of skeletal muscle cells play a particularly prominent role as there is

a high demand for ATP due to the intermittent contractions of the muscle sarcolema[48].

Muscle mitochondrial play another important metabolic role by fine-tuning their oxidative

metabolism to the ambient nutrient status[49]. To illustrate this point, during times of

starvation, the muscle will use lipids as their main source of oxidative fuel. On the other hand,

in the fed state, carbohydrates will replace lipids as the primary substrate for mitochondrial

metabolism[49]. Muscles are one of the largest contributors to total body mass and they are

the most significant tissue involved in insulin-stimulated glucose uptake[50]. Taken together,

any defect in the muscle mitochondria can conceivably affect whole-body metabolic

homeostasis.

The evidence linking mitochondrial dysfunction with diabetes began with observations in

humans with insulin resistance who exhibited an associated decreased activity of citrate

synthase, carnitine palmitoylotransferase 1, malate dehydrogenase, and cytochrome

oxidase[51]. From this, one can discern a possible mechanism whereby impaired mitochondria

can lead to dysfunctional processing of fatty acids and carbohydrates. In cases of obesity, the

increased amount of fat can cause a backlog of mitochondrial oxidation leading to an

accumulation of products such as diacylglycerols, ceramides, fatty acyl coenzyme A, and ROS,

which have all shown to be involved in insulin resistance and impaired insulin action[52-54].

8

Figure3: Role of mitochondria and insulin resistance. During insulin resistance, FFA availability is increased, which raises triglyceride storage and intracellular concentrations of lipid metabolites (DAG, ceramides, LCA-CoA). DAG and ceramides induce impairment of the insulin signaling pathway via activation of inflammatory messengers (for example, PKC-delta), which leads to inhibitory serine phosphorylation of IRS. Glucose transport and phosporylation is reduced. Stimulation of PGC-1a and PGC-1b, the main regulators of mitochondrial biogenesis and fatty acid oxidation, is induced by insulin in skeletal muscle. FFA activate PPAR-gamma and PPAR-delta. Stimulation of oxidative capacity, mitochondrial biogenesis, and mitochondrial lipid uptake is impaired in the insulin-resistant state. Thus, whole-body lipid oxidation decreases in humans with obesity and insulin resistance as a result of impaired mitochondrial plasticity. Increased lipid availability may also induce uncoupling of the respiratory chain; reduced oxidation of glycolytic substrates, which uncouples fatty acid oxidation rates from TCA cycle rates; and metabolic inflexibility. Figure adapted from Szendroedi, J., Phielix, E., and Roden, M. Nat. Rev. Endocrinol. 8, 92-103 (2012).

This can lead to reduced ATP production and subsequent reduction in the ability of the cell to

perform energy-dependent metabolic functions. The reasons for these mitochondrial

deficiencies could reflect a decrease in mitochondrial mass, altered intrinsic activity, or a

combination of both. Several lines of evidence suggest human insulin resistance is linked to

9

impaired mitochondrial function in vivo[55, 56]. To illustrate this notion, electron microscopy

studies of human skeletal muscle showed a significantly reduced mitochondrial mass in patients

with T2D[57]. While this indicates that decreased mitochondrial density may play a causal

factor in diabetes, studies have also shown that intrinsic mitochondrial respiration was

impaired in humans with T2D[58-60]. Global gene analysis has also revealed a reduction

(approximately 20-30%) in the levels of several OXPHOS pathway genes in diabetic humans with

T2D[61]. Interestingly, reduced OXPHOS expression/function has been observed in response to

obesity[62], high-fat diets[63], and reduced physical activity[64] although the exact molecular

mechanisms remain unclear.

The role of mitochondria in other metabolically relevant tissues e.g. adipose also mediates

aspects of whole-body metabolic homeostasis. Brown adipose tissue, for example, exhibits high

flux of fuels through their mitochondria leading to the generation of heat in a process called

thermogenesis[65]. The proton motive force in these mitochondria is mainly dissipated by the

uncoupling protein, UCP1, leading high levels of fatty acid beta-oxidation resulting increased

themogenic activity and whole-body energy expenditure[66]. While the extent of brown

adipose tissue activity influence on whole-body metabolism is not fully understood, studies

show that these activities are reduced in obesity[67, 68]. Conversely, white adipose tissue

(WAT) contains low amounts of mitochondria and their influence on whole-body metabolism is

speculative. What’s known is that WAT can sense nutritional and hormonal signals and relays

this to its mitochondria which can then metabolize fatty acids and carbohydrate or conversely,

store these fuels as triglycerides[69]. Improper utilization of glucose and lipid in WAT can lead

to altered hormonal levels affecting whole-body metabolism. It has been shown that

10

mitochondrial content in white adipose tissue is decreased in cases of obesity with concurrent

insulin resistance[70]. It is conceivable that insulin resistance induced by dysfunctional

mitochondrial WAT can indirectly impact on beta cells by precipitating hyperglycemia and

subsequently exhausting beta cell insulin production and release.

As mentioned earlier, the mitochondria of beta cells play a particularly important role in

glucose homeostasis by virtue of the fact that metabolism in this cell type is coupled to insulin

secretion. The beta cell mitochondria produce ATP, ROS, and products of anaplerosis which are

required for normal insulin secretion[45, 71, 72]. The effects of mitochondrial dysfunction are

exemplified by beta cell-specific mitochondrial transcription factor A (Tfam) knockout mice

which presented with a form of diabetes called mitochondrial diabetes, which is believed to

account for up to 1% of all diabetes cases[73]. One proposed mechanism by which

mitochondrial dysfunction leads to beta cell failure is via chronically increased levels of ROS

which eventually induces beta cell death[74, 75]. The formation of mitochondrial ROS is highly

regulated by the uncoupling protein, UCP2, which acts to safeguard the beta cell from

excessively high levels of ROS produced by the activity of the ETC[76]. UCP2 achieves this by

transporting protons back into the matrix, thereby diminishing the proton gradient and

negatively affecting ROS production[77]. This major influence and dichotomy of UCP2 has

prompted several studies investigating its role in diabetes. Though reports are conflicting, it is

undeniable that improper level of UCP2 activity is detrimental to the beta cell[77]. It was shown

in 2006 that UCP2 can be regulated at the mRNA level by SIRT1, a protein that binds to the

promoter of Ucp2 and represses its transcription[78]. The idea that SIRT1 is a regulator of

11

mitochondrial dynamics has become an attractive topic, and one that has warranted further

investigation.

1.2 Introduction to SIRT1

1.2.1 Sirtuin Family

Transcriptional repression in yeast is mediated by a family of SIR proteins which act to silence

regions of the genome by deacetylation of histones[79]. In doing so, the histone can fold into a

more compact state, rendering the nucleosome inaccessible to transcriptional machinery[80].

The biological processes regulated by SIR2 include senescence/longevity[81], DNA repair[82,

83], and stress responses to heat and starvation[84, 85]. Caloric restriction was shown to

strongly impact aging by slowing it down in yeast and it demonstrated that SIR2 was required

for this robust method of anti-aging to occur [86, 87]. The importance SIR2 is further exhibited

by its high level of conservation of structure and function from archaea to humans[88]. The

mammalian homolog of SIR2 is SIRT1 which stands for (silent mating type information

regulation 2 homolog) 1 shares the greatest degree of sequence identity to SIR2[89] compared

with SIRT2-7[90]. SIRT1, 6, and 7 are localized primarily in the nucleus, SIRT3, 4, and 5 in the

mitochondria and SIRT2 in the cytosol[91]. SIRT1 is the most well-studied of the sirtuins and will

be the focus of this thesis. Although SIRT1 has been observed in the cytosol[92], the extent of

its localization depends on nuclear import/export signals[93]. SIRT1 functions as a deacetylase

which is dependent on the co-factor NAD+ for its activity[94]. As a class III histone deacetylase,

SIRT1 targets include histone H1, H3, and H4 which promotes the formation of

heterochromatin thereby regulating a significant array of biological functions, similar to

12

SIR2[95, 96]. Studies in yeast have revealed that SIR2 contains a Rossmann fold and a smaller

zinc-containing domain which forms a pocket for NAD+ to bind[97]. This pocket also

accommodates the entry of an acetyl-lysine side-chain of various SIRT1 targets where SIRT1 will

catalyze the transfer of the acetyl group onto the ADP-ribose structure of NAD+

forming

nicotinamide (NAM), O-acetyl-ADP-ribose and the deacetylated substrate[98] illustrated by:

NAD+

+ Acetyl-lysine NAM + O-acetyl-ADP-ribose + deacetylated substrate

The levels of SIRT1 activity are dependent on cellular [NAD+]/[NADH] ratios which implicate

SIRT1’s role in regulating metabolism. Moreover, the level of NAD+

is governed by cellular

bioenergetics which ascribes an important role for SIRT1; an energy sensor that allows SIRT1 to

Figure 4: Schematic representation of SIRT1-mediated substrate deacetylation. The acetyl group from the substrate is transferred to the highlighted oxygen group of the ribose ring to yield 2’-O-Acetyl-ADP Ribose and nicotinamide (NAM). Adapted from Guarente, L. N. Engl. J. Med. 364, 2235-44 (2011).

13

assess the energetic status of the organism[99]. SIRT1’s regulation of metabolism goes beyond

its NAD+

1.2.2 SIRT1 Biological Function and Targets

-requirement - it has been increasingly clear that SIRT1 targets span beyond histones to

include transcription factors such as forkhead box O1 (FOXO1), PGC1a, and nuclear factor kappa

B(NFkB)[100]. Regulation of targets such as these can influence a wide range of physiological

processes.

SIRT1 is ubiquitously expressed throughout the body in a variety of organs such as the

heart, liver, pancreas, muscle, adipose tissue, intestinal cells, and throughout the brain[101,

102]. It has also been observed to a lesser extent in the lungs, spleen, thymus, and sex

organs[102]. This gives rise to a number of different cell-specific effects such as cell-cycle

regulation, circadian rhythms, aging, and metabolism. The importance of SIRT1 is aptly

illustrated by SIRT1 whole-body knockout mice who frequently die within a week after

birth[103]. The ones that survived exhibited a significant hindrance to thrive including

exencephaly, smaller body size, lung, cardiac, and optical defects[103]. The reasons for these

defects are plentiful due to SIRT1’s vast array of substrates.

One of the early targets that SIRT1 was discovered to deacetylate was p53[104]. p53 is pivotal

in the DNA damage response, halting the cell cycle to allow for repair of the genome, and

apoptosis[105]. Upon successful completion of these processes, p53’s imposed blockage must

be uplifted so that the cell may continue with its normal functions or p53 can induce apoptosis

if the cell is damaged beyond repair. SIRT1-mediated deacetylation of lysine residue 382 of p53

inhibits p53’s actions and may serve to restore pre-damaged cellular activity[104]. However, if

14

programmed cell death is the cell’s fate, and if apoptosis is overactive, it can lead to

pathological states[106]. SIRT1 is thus considered to be an anti-apoptotic factor. However, its

role in apoptosis may also be due to its ability to deacetylate Ku70, another pro-apoptotic

factor induced by stresses[107]. SIRT1 also deacetylates several more factors involved in cell

cycle and DNA repair, all of which contribute to the notion that SIRT1 is a tumor suppressor

[108-112]. Its link with age-related diseases has been intensely investigated in the past several

years. The Forkhead box group O (FoxO) family of proteins are transcription factors that are

deacetylated by SIRT1 to increase their transcriptional activity in the nucleus[113]. FOXO3

upregulates genes involved in the apoptosis-induced stress[113] while FOXO1 plays a different

role by regulating the expression of metabolic and behavioural genes[114, 115].

An early indication that SIRT1 is involved in metabolism came from Picard et al. who showed

that SIRT1 repressed PPAR-gamma’s actions on downstream genes thereby resulting in

increased fat mobilization[116]. 3T3-L1 cells overexpressing SIRT1 exhibited reduced

adipogenesis and in vivo, mice that were fasted overnight exhibited increased SIRT1 expression

which increased lipolysis and fat mobilization thereby reducing fat mass[116]. This study began

to shed light on the link between the largely unknown interactions of diet, fat, and aging, and

offered a possible mechanism to counter the adversity associated with age-related declines in

health.

Beyond its regulation of PPAR-gamma, SIRT1 influences fat metabolism through a number of

targets involved in hepatic lipid homeostasis[117]. Improper lipid metabolism can result in

diseases such as hepatic steatosis, non-alcoholic fatty liver disease (NAFLD), hepatic

inflammation and insulin resistance, and liver failure[118]. Purushotham et al. showed that

15

SIRT1 interacts with Peroxisome proliferators-activated receptor alpha (PPARa) to positively

regulate its transcriptional actions. When SIRT1 is deleted specifically in the liver, PPARa

signaling is disrupted and downstream beta-oxidation becomes impaired, leading eventually to

hepatic steatosis[119]. SIRT1 also deacetylates and activates LXRs to regulate hepatic fat and

cholesterol metabolism[120], which is related in part to Sterol regulatory element-binding

protein (SREBP) 1c[121], another target of SIRT1. SIRT1 can transmit fasting cues to inhibit

SREBP1c activity, thereby reducing fat storage[122]. This is just a short yet important panel of

studies that have highlighted SIRT1’s role in adipose and hepatic lipid metabolism. However,

the liver is also prone to dysregulated glucose production in disease. This raises the question as

to whether or not SIRT1 is also an important factor in controlling glucose production.

A short time after Picard et al’s discovery of PPAR-gamma regulation by SIRT1, the PPAR-

gamma coactivator-1alpha (PGC-1a) was shown to upregulate gluconeogenic gene expression

during starvation in an HNF4a- and SIRT1-dependent manner[123]. This group demonstrated

that SIRT1 interacts and deacetylates PGC-1a to facilitate HNF4a’s transcriptional activity[123].

Importantly, SIRT1 in this context acts as a nutrient sensor to maintain sufficient blood glucose

levels to supply vital organs such as the brain with nurients[124]. Conversely, fasting causes

glucagon to activate the gluconeogenic response in the liver by dephosphorylation of CREB

regulated transcription coactivator2 (CRTC2/TORC2)[114]. The acetyl transferase P300 is also

activated by glucagon which acetylates CRTC2 to enhance its activity. SIRT1 was shown to

remove this acetyl group causing downregulation of the gluconeogenic genes[114]. These

examples illustrate SIRT1’s reciprocal roles in the liver depends on factors through which it acts.

This dichotomy of SIRT1 may act to fine-tune glucose production during periods of nutrient

16

fluctuations where prolonged starvation may shift the balance towards a modestly elevated

glucose production. While these examples demonstrate SIRT1’s regulation over hepatic glucose

output, this aberrant process can arise due to insulin resistance in the liver, adipose and/or

muscle.

Figure 5: Schematic representation of selected SIRT1 targets. Red targets are activated and blue targets are deactivated by SIRT1. Circled targets are indirectly modulated by SIRT1 i.e. not via deacetylation.

The liver, fat, and muscle are prone to developing insulin resistance in the context of

obesity and inflammation[125]. SIRT1 was shown to be downregulated in insulin-resistant

tissues, which suggests that SIRT1 is an important factor is maintaining insulin-sensitivity[126-

17

128]. For example, genetic deletion of Sirt1 in the liver resulted in hepatic insulin resistance and

excessive glucose production. Interestingly, it also increased ROS production. The exact causes

of insulin resistance are not fully understood however, there is strong evidence that

inflammation and obesity are predominant factors in this disease process. Excessive fat in the

muscle and liver of obese mice can interfere with insulin signaling although interestingly, a

hormone released from adipocytes called adiponectin can travel to these tissues and reduce

intracellular triglyceride content by promoting fatty-acid consumption and energy

utilization[129]. Adiponectin expression is down-regulated in diabetes, and studies have

demonstrated SIRT1 increases adiponectin transcription by activating FOXO1[130]. To further

this point, caloric restriction has been shown to induce high levels of plasma adiponectin in

rats[131], an intervention that also increases SIRT1 expression[132, 133]. Similarly, the

accumulation of WAT as a function of age contributes to the progression of metabolic

syndromes including insulin resistance.

From these examples, we can see that obesity is causally implicated in the development of

insulin resistance. Increased fat also plays an indirect role by contributing to a low-grade

chronic inflammation which underlies the pathogenesis of insulin resistance[134]. These

reports are supported by studies that show anti-inflammatory agents can rescue insulin

sensitivity[135]. Genetic manipulation in mice to create a modest increase in IKK-beta

expression induces NFkappaB activity to similar levels seen in obesity[134]. NFkappaB acts as a

master regulator of proinflammatory cytokines such as IL-6 and TNf-alpha which can signal into

the cell to activate jun-N-terminal kinase 1 (JNK1)[136]. JNK1 phosphorylation of serine residues

on IRS-1 can interfere with its ability to be induced by insulin-stimulated tyrosine kinase

18

phosphorylation[137], offering a plausible link between inflammation and insulin resistance.

Experiments have demonstrated that SIRT1 activation can downregulate inflammatory gene

expression with concurrent increases in insulin signaling[138]. A potential explanation for this is

by way of SIRT1 deacetylation of NF-kappaB to downregulate its transcriptional activity[139].

This is logical because many of the targets under NFkappaB’s control are inflammatory

genes[140].

Figure 6: TNFa ligand binding to the TNF Receptor leads to activation of JNK1. JNK1 may potentially impair insulin signaling by phosphorylation of IRS1.

While these studies are relatively new and still much more work needs to be done to

uncover SIRT1’s role in mediating insulin sensitivity, it is quite clear from the observations

above that SIRT1 is a salient protective factor against insulin resistance. Though, in the context

19

of glucose homeostasis, arguably the most important cell-type to consider is the insulin-

releasing pancreatic beta cell.

1.2.3 SIRT1 and the Pancreatic Beta Cell

Studies in C. Elegans has shown that the longevity effects of Sir2 depends on the FOXO

transcription factor family member, Daf-16 [141, 142]. Furthermore, FOXO1 is intimately

involved in metabolism and insulin signaling[143, 144], therefore, it was logical to investigate

whether SIRT1 could interact with FOXO1 in the beta cell. In 2002, Kitamura et al. showed that

beta cell failure induced by Irs-2 deletion was restored by reduced expression of FOXO1, which

resulted in increased beta cell proliferation[145]. The mechanism for this was through the

mutually exclusive nuclear actions of FOXO1 and PDX1, of which the latter acts to upregulate

genes to enhance beta cell mass. While FOXO1 may seem detrimental in this light, FOXO1 plays

a protective role in the beta cell when subjected to oxidative stress. Beta-TC3 cells treated with

hydrogen peroxide causes translocation of FOXO1 to the specific nuclear subdomains to

increase NeuroD and MafA transcriptional activity[146]. NeuroD and MafA are both known to

increase insulin biosynthesis as well as maintaining the proper function of beta cells[147-149].

The transcriptional co-activation of FOXO1 was dependent on SIRT1-mediated deacetylation,

which also caused more rapid degradation of FOXO1 to switch off its activity. These examples

show that SIRT1 can exert protective effects against oxidative stresses and insulin signaling

impairment on the beta cells through the target FOXO1.

20

Interestingly, more recent studies conducted by Bastien-Dionne et al. showed in pancreatic

beta cells that GLP-1 inhibited Sirt1 expression. Because SIRT1 deacetylates and activates

FOXO1, in this context, GLP-1 acted to inhibit FOXO1 actions through SIRT1 to increase beta cell

expansion[150]. These proliferative effects on the beta cell were abolished by SIRT1

overexpression which lead the investigators to conclude that SIRT1 is a negative regulator of

beta cell growth.

Anti-proliferation can be a mechanism to limit beta cell expansion in diabetes however, beta

cells could similarly be undergoing cell death. Due to the firm link between obesity,

inflammation and diabetes, it could be the case that inflammation not only precipitates insulin

resistance but inflammatory mediators can induce cytotoxicity. A growing number of studies

have demonstrated the impact of cytokines on beta cell dysfunction and death, raising the

point that diabetes is increasingly becoming recognized as a disease of inflammation [151-153].

Amongst the most significant targets of cytokines is NFkappaB which upon activation, will

dissociate from IkappaB, (a regulatory protein that sequesters NFkappaB in the cytosol), and

translocate into the nucleus where it will increase the expression of pro-inflammatory

cytokines, adhesion molecules, chemokines, iNOS, cyclooxygenases (COXs) and matrix

metalloproteinases[154]. The net effect of NFkappaB signaling is to increase immune cells at

the site of inflammation to rid the body of the pathogen and/or insult. Overexpression of SIRT1

in rat islet cells (RIN) showed significantly decreased NFkappaB activity when stimulated with

cytokines[155]. Cytokine-induced cell death was achieved by treatment with IL-1beta and IFN-

gamma, which killed more than half of the RIN cells by way of nitric oxide (NO) production.

21

Increased SIRT1 was able to mitigate NO levels by repressing iNOS expression which ultimately

decreased beta cell death[155].

Beta cell death could similarly be caused by treatment with streptozotocin (STZ) which

specifically targets and kills beta cells[156]. Studies using mice with a mutation called WldS

(Wallerian generation slow) has the effect of increasing NAD levels thereby increasing SIRT1

activity. STZ treatment in WldS+/+

Perhaps the most striking example of SIRT1’s involvement in diabetes comes from a recent

study that identified a family of five, of which four members developed type 1 diabetes at ages

7, 12, 15, and 26. Multiple sequencing methods led to the discovery of a single amino acid

substitution in exon 1 of Sirt1 of these individuals, leucine-to-proline, which was associated

with overproduction of cytotoxic factors; cytokines, chemokines, and NO[158]. Of the many

genes associated with the development of diabetes, Sirt1 is now amongst the very few which

can lead to a monogenic form of diabetes[158], others include AIRE[159] and FOXP3[160].

Intriguingly, recent evidence has implicated SIRT1 as a regulator of FOXP3+ regulatory T

cells[161].

mice exhibited significantly lower hyperglycemia and

increased insulin transcription compared to controls suggesting that SIRT1 in involved in

increasing insulin synthesis and protected animals from STZ-induced diabetes[157].

These examples illustrate the importance of SIRT1 in beta-cells proliferation and death.

However, it is known that in diabetes, there are functional defects of the beta-cell machinery

that render it unable to compensate for hyperglycemia[162-165]. These events can be

independent of changes to beta cell mass, which tip the balance between pre-diabetes and

22

frank diabetes. Understanding how the function of beta cells becomes dysfunctional and result

in reduced insulin secretion is crucial information that is still incompletely understood.

1.2.4 SIRT1 and Insulin Secretion

One year after the discovery of SIRT1 being directly involved in metabolism by several

groups, Moynihan et al. published a paper that explored the effects of overexpressing Sirt1

cDNA under the human insulin promoter to drive expression specifically in mouse pancreatic

beta cells (BESTO)[166]. This was of great interest as beta cells secrete insulin in response to

nutrients, namely glucose. This group showed that in their BESTO mice, Sirt1 was

overexpressed 16-18-fold and when challenged with a glucose tolerance test, they exhibited

improved glucose disposal with an associated increase in insulin secretion. The enhanced

insulin secretion was recapitulated in vitro in isolated BESTO islets stimulated with 25mM

glucose. Similarly in the study using WldS+/+ mice, only those mice with two alleles of Sirt1 intact

showed enhanced insulin secretion compared to WldS+/+Sirt-/-

To determine the underlying mechanisms for this phenotype, Moynihan et al. performed

microarray on Sir2 overexpressing MIN6 cells which among the differentially expressed genes,

Ucp2 was noteworthy. Like Ucp1, Ucp2 also acts to dissipate the proton gradient. However, in

the context of the beta cell, the function is to presumably prevent toxic levels of ROS

accumulation[167]. With Ucp2 downregulated by SIRT1 over-expression, ATP concentration

was enhanced, which could explain the boosted insulin secretion[166]. Interestingly, a much

[157]mice reinforcing the idea

that SIRT1 is a positive mediator of insulin release.

23

smaller increased dose of SIRT1 (approximately two-fold) does not confer the same boost in

insulin secretion upon glucose challenge[168]. However, this modest increase in SIRT1 primed

mice for metabolic adaptions against insulin resistance, reduced energy expenditure and

increased overall metabolic efficiency.

These findings prompted the corollary study whereby Bordone et al. deleted Sirt1 in mice to

investigate the effects on the pancreatic beta cell. As expected, Sirt1 knockout mice showed

lower plasma insulin levels, ad libitum, and after glucose injection[78]. In line with Moynihan et

al’s observations, Ucp2 transcript and UCP2 protein were increased in Sirt1 knockdown cells

and Sirt1 knockout mice[78, 166]. The mechanism of SIRT1’s repression of Ucp2 was

investigated by means of chromatin immunoprecipitation which showed that SIRT1 bound

directly to the Ucp2 promoter to decrease its expression[78]. Together, these findings highlight

a plausible pathway by which SIRT1 increases insulin secretion. However, because of the large

number of SIRT1 substrates, it’s unlikely that Ucp2 is the only target that SIRT1 is regulating in

the pancreatic beta cell. Furthermore, beta cell-specific UCP2 knockout mice (UCP2BKO) are

more glucose intolerant with no in differences insulin secretion during OGTT[169]. Robson-

Doucette et al. found that the UCP2 deletion increases glucagon secretion suggesting that the

alpha cells may be more pertinent in the context of UCP2 and diabetes[169]. Importantly, the

data reinforce the idea that SIRT1 is not mediating its effects on insulin secretion solely through

UCP2 repression, therefore warranting further investigation of the role of SIRT1 in the beta cell.

24

2 Rationale and Hypothesis

Rescuing insulin secretion is of great therapeutic interest for individuals living with diabetes.

What begins as modest fasting plasma glucose can escalate to drastically toxic levels where

insulin release is unable to compensate due to gradual decline in beta cell function.

Tremendous efforts have been put forth over the past decades to understand how the beta cell

functions in the normal setting and how it becomes dysfunctional in diabetes. Although many

molecules and mechanisms have been identified, the glucose-induced insulin secretory

pathway still eludes our complete understanding. In the last decade, studies have identified the

NAD+-dependent deacetylase, SIRT1, as a major factor in metabolism and whole-body glucose

homeostasis by helping to maintain insulin sensitivity and enhancing insulin secretion. The

former was achieved through a number of mechanisms involving many SIRT1 targets which

regulate processes such as adiponectin release, inflammation, and insulin signaling. Enhanced

insulin secretion on the other hand was mainly attributed to decreased Ucp2 expression by

SIRT1. While the idea that SIRT1 is regulating mitochondrial bioenergetics through Ucp2 to

affect insulin secretion is attractive, given the large number of targets that converge on

mitochondrial function, it is likely that Ucp2 does not offer the entire picture. Furthermore

there have been overexpression studies in whole-body and beta cells in mice, previous studies

have used global Sirt1 deletion from birth rendering the effects on glucose homeostasis difficult

to interpret. Although these studies showed important proof of principle, the confounding

effects of embryonic whole-body knockouts have their limitations. Therefore, studying SIRT1

25

specifically in the pancreatic beta cells in a spatially and temporally controlled manner is

imperative. I hypothesize that deletion of Sirt1 specifically in pancreatic beta cells will impair

insulin secretion by perturbing mitochondrial bioenergetics and precipitate a glucose intolerant

phenotype upon glucose challenge. These affects can be directly attributable to altered events

from the pancreatic beta cell and not other peripheral and/or endocrine cells. I believe by

studying the beta cell in this context will reveal molecular changes beyond mitochondrial

uncoupling protein 2 that are responsible for changes that occur in diabetes. This novel

information can then be used to design therapeutics to treat individuals living with diabetes.

26

3 Materials and Methods

3.1 Experimental SIRT1 Inactivation Models

3.1.1 Sirt1BKO Mice

Pdx1creER mice (created by the laboratory of Dr. Doug Melton, Harvard University,

Cambridge, MA, USA) were crossed with homozygous floxed Sirt1 exon 4, which encodes the

catalytic domain of SIRT1 (a kind gift from Dr. F. Alt., Harvard, Boston, MA, USA) mice.

Figure 7: Schematic of tamoxifen-inducible CreER Lox system. Step 1: Floxed Sirt1 Exon 4 (Ex4) mice crossed with Pdx1CreER mice yielding mice with CreER protein expressed specifically in pancreatic beta cells. Step 2: administration of tamoxifen will bind to CreER protein that has been sequested in the cytosol to induce translocation into the nucleus where Sirt1 Ex4 alleles can then be recombined.

27

Tamoxifen (125mg/kg) or corn oil vehicle was injected intraperitoneally in 9- to 10-week-old

male Pdx1CreER:floxSirt1 mice to produce pancreatic beta cell-specific Sirt1 knockout

(Sirt1BKO) or control mice, respectively. Experiments were performed 5-10 days after the last

injection (injections were administered on alternating days for a total of three injections). Sirt1

exon 4 deletion was validated by PCR and knockdown using primers designed to span exon 4

was validated via qPCR. Truncated SIRT1 protein was confirmed by means of western blot. All

the protocols were approved by the animal care committee of the University of Toronto.

3.1.2 SIRT1KD MIN6 Cells

Insulinoma pancreatic beta cell lines created by targeted expression of simian virus 40 T

antigen gene in mice (MIN6 cells)[170] (a kind gift from Dr. S. Seino, Chiba University, Chiba,

Japan) were seeded at 40–50% confluency onto 24-well plates and transfected with 300 nmol/l

Sirt1 short interfering RNA (SirT1 SMARTpool siRNA; Dharmacon, Thermo Scientific, Waltham,

MA, USA) (SIRT1KD) or scrambled siRNA (control) (Si-Scram) using Lipofectamine 2000

(Invitrogen, Burlington, ON, Canada) according to the manufacturer’s instructions. Cells were

maintained in DMEM containing 15% FBS, 100U/ml P/S and 1.7μL beta-mercaptoethanol per

500ml. After 48 h transfection, cells were used for analysis. Sirt1 knockdown was validated by

means of qPCR and protein knockdown was confirmed by western blot.

3.2 Glucose and Insulin Tolerance Test

Glucose tolerance test (GTT): following a 14 h fast, mice were mice were given 2 g/kg

glucose by oral gavage and blood glucose was measured from tail vein using a glucometer

(Bayer, Tarrytown, NY, USA) at 0, 10, 20, 30, 60, and 120 min. Blood was collected at 0, 10, 20,

28

30, 60, and 120 min in EDTA-coated microcuvettes for plasma insulin measurements. Plasma

insuliln was measured by ELISA (Alpco, Salem, NH, USA).

Insulin tolerance test (ITT): following a 4 h fast, mice were given an intraperitoneal injection of

1 U/kg insulin. Blood samples were taken at 0, 10, 20, 30, 60, and 120 min from the tail vein

using a glucometer (Bayer, Levekusen, Germany) as previously described[169].

3.3 Islet Isolation and Dispersion

Mice were anesthetized with an intra-peritoneal injection of 250 mg/kg

tribromoethanol. The pancreas was perfused via the common bile duct with collagenase type-V

(0.8 mg/ml) in RPMI-1640 media supplemented with 2% BSA and 1% penicillin/streptomycin.

Pancreata were digested at 37o

C for 15 min and islets were hand-picked and cultured in RPMI-

1640 media supplemented with 10% FBS and 1% penicillin/streptomycin as previously

described[171]. Dispersion: Isolated islets were washed in PBS supplemented with 2 mmol/l

EGTA and dispersed onto glass coverslips using Accutase (Invitrogen) for 5 min at 37 ⁰C.

3.4 Glucose – and KIC-stimulated Insulin Secretion

Isolated islets or MIN6 cells were equilibrated in Kreb’s Ringer Buffer (KRB) (mmol/l:

NaCl 115, KCl 5, NaHCO3 24, CaCl2 2.5, MgCl2 1, HEPES 10, glucose 2, BSA 0.1%) for 1 h and

stimulated with the indicated concentration of glucose or alpha-ketoisocaproate (KIC) for 1 h.

The supernatant fraction was collected for insulin measurement by RIA (Linco Research,

Millipore, St Charles, MO, USA) as previously described[171] or HTRF following the

manufacturer’s protocol (Cisbio, Bedford, MA, USA). For Resveratrol (25umol/l) and/or Sirtinol-

29

supplemented (25umol/l) GSIS experiments, islets or cells were incubated in their respective

media 48 h prior.

3.5 Western Blot

Isolated mouse islets (100–150) were lysed in RIPA buffer (Cell Signaling, Danvers, MA,

USA) containing protease inhibitor cocktail (Roche, Mississauga, ON, Canada). Lysates were

spun at 11,200 g and the supernatant fraction was electrophoresed on a 4–15% SDS-PAGE

gradient gel (BioRad, Mississauga, ON, Canada) and then transferred onto a polyvinylidene

difluoride (PVDF) membrane using a Turbo Blotter (BioRad). The membrane was probed with

anti-SIRT1 antibody at 1:1000 (Cell Signaling), anti-Gβ at 1:1000 (Santa Cruz Biotechnology,

Dallas, TX, USA) and/or anti-acetyl lysine at 1:1000 (Cell Signaling) and imaged using Kodak

Imager 4000pro (Carestream, Rochester, NY, USA) as previously described[172].

3.6 Polymerase Chain Reaction and Microarray

Total RNA was extracted from isolated mouse islets or MIN6 cells with an RNeasy Mini Kit

(Qiagen, Germantown, MD, USA) and converted to cDNA using SuperScript II reverse transcriptase

(Sigma-Aldrich, St Louis, MO, USA). Real-time PCR was performed as previously described [171, 173] on

the Dual Block DNA Engine Thermal Cycler (MJ Research, Waltham, MA, USA) as previously

described[174]. Primers were designed using Primer Blast (NCBI, Bethesda, ML, USA). 10 ng of cDNA per

well was used as the template for quantitative PCR amplification or run on a 1% agarose gel for gel

electrophoresis PCR. The qPCR protocol was as follows: heat activation of polymerase at 95°Cfor 3 min,

followed by 40 cycles of 95°C for 10 s, 65°C for 15 s, and 72°C for 20 s. Readings were carried out on an

30

ABI Prism7900HT Sequence Detection System (Applied Biosystems, USA) and compared against a

standard curve created from mouse genomic DNA by serial dilutions. Data were normalized to mouse β-

actin mRNA.

Microarray was performed at the University Health Network Microarray Centre (Toronto, ON,

Canada) using the Affymetrix Mouse 430_2.0 Gene Chip on total RNA extracted from SIRT1KD

and scrambled control cells. Data were annotated by selecting only those genes whose

expression changed significantly at least 1.3-fold (up and down) and analyzed by PANTHER

Gene Ontology™ (http://www.pantherdb.org/) to classify genes into biological processes as

previously described[175] .

3.7 Immunohistochemistry

Sirt1BKO and control mice were sacrificed and their pancreata were removed surgically by hand,

weighed and fixed in 10% neutral buffered formalin for 48 h. Sample preparation was performed by the

Department of Pathology, Toronto General Hospital. Samples were embedded in paraffin and

histological sections were prepared from each pancreas and mounted on slides. Sections were labelled

with rabbit polyclonal anti-insulin (1:200 dilution) or rabbit polyclonal anti-glucagon (1:150 dilution) as

previously described[176]. Slides were digitized on a bright-field scanner at 20 times magnification

Images of each section were acquired using Aperio Imagescope version 11.0.2.725 (Aperio Technologies,

Vista, CA, USA) at 40× magnification. The beta and alpha cell area was calculated by positive pixel

analysis using Aperio Imagescope software.

31

3.8 Transmission Electron Microscopy

Freshly isolated islets were pelletted by centrifugation and fixed with 2.5% glutaraldehyde in

0.1M cacodylate buffer, pH 7.4, for 1 h at room temperature. Sample preparation was performed by the

Microscopy Imaging Lab, University of Toronto. After rinsing with cacodylate buffer, pellets were post-

fixed in 1% osmium tetraoxide for 2 h, dehydrated in a graded series of ethanol and then embedded in

Epon. 60-80 nm thick sections were mounted on copper grids, and stained with uranyl acetate and lead

citrate. Samples were observed under a Philips CM100 electron microscope operating at 75 kV and

images were acquired digitally using a Kodak 1.6 Megaplus camera system operated using AMT software

(Advanced Microscopy Techniques Corporation) as previously described[177]. Mitochondrial area was

quantified using ImageJ software version 1.46r (NIH, Bethesda ML, USA).

3.9 Calcium Imaging and Mitochondrial Membrane Potential

Changes in intracellular calcium concentrations were assessed using Fura-2 AM (Invitrogen) in

dispersed islet cells as previously described [175]. Cells were loaded with 2 μmol/l Fura-2 AM dye for 50

min at 37°C. Cells were perifused with bath solution at 1 ml/min at 37°C. Experiments were performed

using an Olympus BX51W1 microscope (Tokyo, Japan) with an ×20/0.95 water immersion objective and

cooled charge-coupled device camera. Measurements were taken using ImageMaster version 3.0

software (Photon Technology International, London, ON, Canada). Analysis was performed using Igor

Pro version 4.0 software (Wavemetrics, Lake Oswego, OR, USA) and normalised to baseline. Only cells

showing response to 30 mmol/l KCl solution were included. Mitochondrial membrane potential (MMP)

was measured as previously described using rhodamine-123 (Invitrogen) [178].

32

3.10 Oxygen Consumption Rate

siRNA-treated MIN6 cells were plated at 50,000 cells per well into XF24 plates (Seahorse

Bioscience, Billerica, MA, USA). Cells were pre-incubated in KRB buffer (0 mmol/l glucose) for 1.5 h at

37°C before loading into an XF24 respirometry machine (Seahorse Bioscience). Oxygen consumption

rates (OCRs) were measured at 0 and 20 mmol/l glucose, 5 µmol/l oligomycin, 50 µmol/l 2,4-

dinitrophenol (DNP) and 10 µmol/l rotenone + myxothiazol in the configuration: (in min) mixing 2,

waiting 2, measuring 3, loop ×4. Raw OCR data was normalised to total RNA (ng). These values were

then expressed as a percentage of the maximal OCR (as measured with DNP treatment).

3.11 ATP Measurements

Isolated islets were equilibrated in 2mmol/l glucose KRB for 1 h prior to stimulation with

2mmol/l or 20mmol/l glucose for 15 min. Islets were then treated with 100ul of 1x ATP extraction buffer

from the StayBrite™ Highly Stable ATP Bioluminescence Assay Kit (Biovision, Milpitas, CA, USA) and

homogenized. Homogenate was spun at 10,000rpm for 2 min and supernatant was collected for ATP

measurements following manufacturer’s protocol.

33

4 Results

4.1 Validation of SIRT1 Inactivation To study SIRT1’s function specifically in adult beta cells, we employed a cell-specific

inducible deletion strategy as previously described [179]. Although expressed in whole

pancreas during the embryonic phase, the Pdx1 promoter is expressed exclusively in adult beta

cells. The intraperitoneal administration of tamoxifen induced deletion of floxed exon 4, which

encodes the catalytic domain of Sirt1. This is demonstrated by the presence of a smaller band

(390 bp) in Sirt1BKO islet mRNA compared with wild-type mice (543 bp) (Fig. 8a). A faint lower

band was observed in the vehicle-injected controls indicating some CreER activity independent

of tamoxifen (Fig. 8a). When measured by qPCR, the level of Sirt1 mRNA in Sirt1BKO mouse

islets was ~70% lower than that in control and wild-type mouse islets (Fig. 8b). SIRT1

inactivation was further confirmed by western blotting. A lower band was observed in Sirt1BKO

islets compared with controls using an antibody recognising the c-terminus of SIRT1 (Fig. 8c).

Recently, CreER expression has been detected in the hypothalamus of transgenic Pdx1-Cre

mice, suggesting that the Pdx1 promoter may be active there [180]. Importantly, hypothalamic

Sirt1 deletion was not observed by PCR or qPCR in Sirt1BKO mice. Sirt1BKO experiments were

run in parallel with Sirt1 siRNA-treated mouse insulinoma pancreatic beta cell line, MIN6,

(SIRT1KD) resulting in greater than 75% knockdown (Fig. 8d). Western blot analysis showed

marked protein knockdown (Fig. 8e). Sirt1 mRNA expression as quantified in non-islet metabolic

34

tissues e.g. muscle, white adipose tissue (WAT), duodenum, and liver. There was no difference

between Sirt1BKO mice and vehicle-controls indicating that the Pdx1 promoter is not active in

the tissues (Fig. 8f). These data show that Sirt1 is inactivated in the beta cells of Sirt1BKO mice

and SIRT1KD MIN6 cells, specifically in mouse pancreatic beta cells in the former.

Figure 8: The inactivation of Sirt1 in Sirt1BKO mice and SIRT1KD MIN6 cells. (a) RT-PCR (A, no treatment; B, corn oil-injected; C, tamoxifen-injected) and (b) qPCR analysis of Sirt1 in islets and hypothalamus (hypo) from floxed (grey), control (white) and Sirt1BKO mice (black bar). (c) Western blot of SIRT1 in Sirt1BKO islets. (d) Decreased Sirt1 mRNA in SIRT1KD MIN6 (black bar) vs scrambled controls (Si-Scram) (white bar). (e) Western blot of SIRT1KD and scrambled controls (Si-Scram). (f)

35

Sirt1 mRNA expression non-islet metabolic tissues, WAT (White Adipose Tissue). n=3–5 for each group of mice. n=3–5 independent experiments run in replicates of six for SIRT1KD MIN6. *p<0.05 vs control and floxed;**p<0.01 vs scrambled controls

4.2 Sirt1BKO Mice are Glucose Intolerant

Since SIRT1 overexpression was shown to enhance insulin secretion, it is logical that

Sirt1 deletion in the beta cell could potentially affect glucose homeostasis and metabolism.

Tamoxifen injections could potentially affect metabolism and body weight therefore weight

was monitored prior to injections (9-weeks-old) and after injections (12-week-old).

Figure 9: Impaired glucose tolerance and decreased insulin secretion in Sirt1BKO mice. (a) Body

36

weight prior to tamoxifen injection (9-week-old) and b) post-injection (12-week-old). (c) Blood glucose and (d) plasma insulin (ad libitum). Sirt1BKO (black bar) and controls (white bar). (e) OGTT (2 g/kg glucose) performed after a 14 h fast in control (square) and Sirt1BKO mice (circle), n=12 for both groups. (e) Plasma insulin levels during OGTT. (f) ipITT test (1U/kg insulin). The blood glucose concentrations were determined at the indicated times. n=10–12 for both groups. *p<0.05

There was no difference between Sirt1BKO and control mice demonstrating that tamoxifen

does not alter the body weight of mice (Fig. 9a, b). Furthermore, food intake (data not shown)

was similar between the two groups. Blood glucose and plasma insulin (ad libitum access to

food and water) were not significantly different between the two groups (Fig. 9c,d). However,

when the Sirt1BKO mice were challenged with glucose after fasting, they showed impaired

glucose tolerance at 20, 30, and 60 min post glucose gavage (Fig. 9e). This was accompanied by

reduced plasma insulin at 10 min after glucose gavage (Fig. 9f). Sirt1BKO mice also had normal

insulin sensitivity during intraperitoneal insulin tolerance tests (ipITT) compared with controls

(Fig. 9g), suggesting that the glucose intolerance was not due to changes in insulin resistance

but rather, due to impaired glucose release into the blood.

4.3 SIRT1 Inactivation Reduces Glucose-stimulated Insulin Secretion

To examine whether the decreased GSIS was a direct effect of Sirt1 inactivation in beta

cells, we isolated islets from Sirt1BKO mice and performed secretion assays. There was no

difference in insulin secretion at low glucose concentration (2 mmol/l) between Sirt1BKO and

control islets (Fig. 10a). However, Sirt1BKO islets secreted significantly less insulin (40%) when

stimulated with 20 mmol/l glucose (Fig. 10a). There was no difference in insulin secretion with

30 mmol/l arginine stimulation, indicating that ion-channel function and cellular depolarisation

are not affected by Sirt1 inactivation. The attenuation of GSIS from Sirt1BKO islets was not

37

caused by diminished insulin content (Fig. 10b) or diminished pancreatic beta cell area (Fig.

10c). Insulin and glucagon staining of pancreatic sections did not reveal any differences in beta

cell or alpha cell morphology, respectively, compared with controls (Fig. 10d–f), indicating that

the glucose intolerance was not due to increased glucagon production.

Figure 10: Decreased insulin secretion in Sirt1BKO islets. (a) Glucose- and glucose+arginine-stimulated insulin secretion from Sirt1BKO (black bar) and control islets (white bar). Glc, glucose; Arg, arginine. (b) Insulin content from GSIS islets and (c) beta and (d) alpha cell area. n=7–15 for each group and each group used 50–220 islets. (e) Insulin staining and (f) glucagon pancreatic cross-section staining. n=3 mice per group with three cross sections per pancreas. *p<0.05

38

4.4 The Effects on Insulin Secretion are Due to Sirt1 Inactivation

In accordance with Sirt1BKO islets, SIRT1KD MIN6 cells also secreted significantly less

insulin when stimulated with 20 mmol/l glucose compared with scrambled siRNA controls (Fig.

11a). This was correlated with decreased SIRT1 activity as measured by increased levels of

acetylated protein in SIRT1KD MIN6 whole-cell lysates compared with controls (Fig. 11b). Wild-

type CD1 islets treated with the SIRT1 activator Resveratrol for 48 h had enhanced GSIS (Fig.

11c).

39

Figure 11: SIRT1 is responsible for changes in insulin secretion. (a) Decreased Glc-stimulated insulin secretion from SIRT1KD MIN6 cells (black bar, SIRT1KD MIN6; white bar, scrambled controls). (b) Acetyl lysine (AcK) western blot of SIRT1KD MIN6 and scrambled controls run in duplicate. (c) Non-treated (white bar), resveratrol-treated (black bar) and resveratrol+10 µmol/l EX527-supplemented (grey bar) GSIS from wild-type CD1 islets. (d) Non-treated (white bar), resveratrol-treated (black bar) and resveratrol+25µmol/l sirtinol-supplemented (grey bar) GSIS from wild-type CD1 islets. (e) Resveratrol-supplemented GSIS in Sirt1BKO (black bar) and control (white bar) islets. Resveratrol concentration: 25 µmol/l and 48 h treatment in all experiments. n=3–15 for each group and each group used 30–220 islets and n=3 independent experiments run in replicates of six for SIRT1KD MIN6.*p<0.05

Accordingly, the SIRT1-specific inhibitors EX527 (10 µmol/l) and Sirtinol (25 µmol/l) prevented

resveratrol from enhancing GSIS (Fig. 11c,d). Importantly, Sirt1BKO islets did not exhibit

enhanced GSIS when treated with 25 µmol/l resveratrol for 48 h (Fig. 11e). These data suggest

that specific inactivation of SIRT1 disrupts GSIS in Sirt1BKO and SIRT1KD MIN6 cells. Conversely

increased SIRT1 activity acutely enhanced insulin secretion.

4.5 Defects in Insulin Secretion are Downstream of Glycolysis

To determine the mechanism by which Sirt1 inactivation impaired GSIS, we examined

several steps of the canonical glucose-induced insulin secretory pathway, beginning with

glycolysis. qPCR analysis did not reveal any significant changes in glycolytic enzyme and Glut2

(also known as Slc2a2) expression, nor was glucose uptake affected by SIRT1 knockdown (data

not shown). To confirm that the defect is downstream of glycolysis, isolated Sirt1BKO and

control islets were subjected to static secretion assays with 2 mmol/l and 20 mmol/l α-

ketoisocaproate (KIC) in place of glucose. KIC is converted to acetyl-CoA, which enters directly

into the TCA cycle to carry out the remaining steps of metabolism [181], effectively bypassing

glycolysis. Sirt1BKO islets secreted significantly less insulin after exposure to 20 mmol/l KIC

40

compared with controls (Fig. 12a). Interestingly, there was a modest yet significant increase in

insulin secretion in Sirt1BKO islets at 2mmol/l KIC. Therefore, the defect in insulin secretion due

to Sirt1 inactivation is downstream of glycolysis.

Figure 12: Reduced alpha-ketoisocaprioic acid-induced insulin secretion in Sirt1BKO mice. Alpha-ketoisocaprioic acid (KIC)-stimulated insulin secretion from control (white bar) and Sirt1BKO islets (black bar). n=140-160 islets from 5-7 mice.

4.6 Sirt1 Knockdown Causes Significant Dysregulation of Metabolic Genes

SIRT1 can directly influence transcription of some genes [78]. Therefore, a global

microarray was performed on SIRT1KD MIN6 cells to gain insight into the cause of impaired

GSIS. Three-hundred-and-forty genes had significantly changed expression (at least 1.3-fold)

compared with scrambled controls. Of the significantly changed genes, 46.5% are involved in

metabolic processes (Fig. 13a), including 37 mitochondria-related genes whose expression was

either up- or downregulated (Fig. 13b). Several components of the electron transport chain,

including Cox7a1 and Cox8a, had increased expression levels (1.32-fold and 1.43-fold,

respectively) relative to controls. The ATP synthase

41

Figure 13: Dysregulation of mitochondria-related genes in SIRT1KD MIN6 cells. (a) Pie chart representation of those genes whose expression changed significantly (either up or down, at least 1.3-fold). (b) Panel of 37 mitochondria-related genes in SIRT1KD MIN6 (black bar) expressed as fold-

42

change from control selected from metabolic group of pie chart. n=3 independent experiments run in triplicate. *p<0.05

subunit, Atp5g1, however, showed a 1.3-fold decrease in mRNA in SIRT1KD cells.

The transcript that displayed the largest change was Pgc1a (also known as Ppargc1a), which

increased 2.4-fold (Fig. 14a). UCP2 has long been held to be responsible for SIRT1’s effects on

insulin secretion; however, our SIRT1KD showed no difference in Ucp2 transcript levels

compared with controls (Fig. 14c). Furthermore, Price et al showed that SIRT1’s beneficial

effects are tied to increased mitochondrial biogenesis [182] but SIRT1KD cells showed no

difference in genes regulating this process (data now shown). Mitochondrial area did not

different significantly between Sirt1BKO islets and controls (Fig 14c-e). Taken together, our

beta cell-specific SIRT1 knockdown model shows that novel mitochondrial respiratory factors,

such as Cox7a1, Cox8a, and Atp5g1, are affected without changes to mitochondrial mass.

Figure 14: Sirt1 Inactivation does not affect mitochondrial mass. (a-b) mRNA levels of Pgc-1a and Ucp2 from control (white bar) and SIRT1KD (black bar) MIN6 cells. n=3-5 for each group run in triplicate. (c) Mitochondrial

43

area calculated from (d-e) electron microscopic images of control and Sirt1BKO islets n=3 for each group. *p<0.05

4.7 SIRT1 Inactivation Impairs Mitochondrial Function

Due to the large cluster of mitochondria-related genes altered by Sirt1 deletion,

mitochondrial function was tested by means of MMP and OCR. Exposure to high glucose (20

mmol/l) concentrations caused decreased hyperpolarization in the mitochondrial matrix of

dispersed Sirt1BKO islet cells compared with controls (Fig. 15a,b). This indicates there is a

reduced proton motive force possibly owing to inefficient respiration. Sodium azide (NaN3)

halts mitochondrial respiration by inhibiting complex IV of the electron transport chain, causing

maximal depolarisation of MMP. There was no difference in NaN3-induced depolarisation

between Sirt1BKO and control cells, excluding dysfunctional channel activity as a possible

reason for decreased hyperpolarisation. Furthermore, SIRT1KD cells showed significantly

reduced OCR under low (0 mmol/l) and high (20 mmol/l) glucose compared with controls (Fig.

14c). Oligomycin is an ATP Synthase inhibitor therefore any oxygen consumption that occurs in

the presence of oligomycin is not coupled to ATP production and is a measure of proton leak.

There was no difference in OCR between Sirt1KD and control MIN6 cells treated with

oligomycin indicating that proton leak was not affected by Sirt1 knockdown (Fig. 15c).

Rotenone + myxothiazol are inhibitors of Complex I and Complex III of the ETC respectively,

which acts to halt respiration. Thus, OCR measured in the presence of these drugs is a measure

of non-mitochondrial oxygen consumption, i.e. cytosolic. There was no difference between

SIRT1KD and controls in this parameter suggesting that SIRT1 does not markedly affect cytosolic

44

oxygen usage in pancreatic beta cells (Fig. 15c). This indicates that the reduced OCR was

primarily due to respiration deficits not involving cytosolic and/or uncoupling processes. These

data highlight a marked impairment of mitochondrial function in SIRT1KD and Sirt1BKO cells.

Figure 15: Sirt1 inactivation impairs mitochondrial hyperpolarization and OCRs in islets from Sirt1BKO and SIRT1KD mice. (a) Representative kinetic traces of the rhodamine-123 (Rhod123) fluorescent signal and (b) summary of MMP changes expressed as per cent change from basal in control (white bar/square) and Sirt1BKO beta cells (black bar/circle). (c) OCR under basal conditions (0 mmol/l glucose) and following exposure to 20 mmol/l glucose, 5 µmol/l oligomycin, 50 µmol/l DNP, and rotenone+myxothiazol (10 µmol/l for both) in SIRT1KD (black circle) and scrambled controls (white

45

square) MIN6. Glc, glucose. n=2 or 3 independent experiments, and each experiment used ~30–100 cells from 3-4 mice per genotype (Sirt1BKO) or n=7–12 (SIRT1KD). *p<0.05

4.8 ATP Production is Impaired by Pharmacological SIRT1 Inhibition

Due to the impaired mitochondrial function in Sirt1BKO and SIRT1KD cells, it was logical

to measure downstream ATP production. MIN6 cells incubated for 15 minutes in low glucose

(0mmol/l) and high glucose (20mmol/l), with and without 25µmol/l Resveratrol

supplementation were lysed and ATP content was quantified. ATP production was enhanced

under both low and high glucose conditions with Resveratrol compared to non-Resveratrol-

treated MIN6 cells (Fig. 16a). These results were recapitulated in isolated CD1 islets incubated

in low and high glucose supplemented with either 25µmol/l Resveratrol or 25µmol/l Sirtinol,

the latter being a SIRT1-specific inhibitor. Resveratrol had the effect of increasing ATP

production while Sirtinol decreased it (Fig. 16b) reinforcing the mitochondrial dysfunction in

Sirt1-inactivated cells. It follows that reduced ATP production should impact on events

downstream such as voltage-gated calcium influx into the cell, which was investigated next.

4.9 Decreased Calcium Influx in Sirt1BKO islet Cells

Recently, it was shown that SIRT1 could influence intracellular calcium accumulation in

certain models of diabetes [183]. Thus, we tested whether SIRT1 could regulate calcium in the

beta cells, as Ca2+ triggers insulin granule release but is also an important cofactor in many

mitochondrial enzymes that stimulate respiration [184-186]. There was no difference in

intracellular calcium ([Ca2+]i) under low glucose (1.0 mmol/l) conditions between Sirt1BKO cells

46

and controls (Fig. 16c). However, when cells were exposed to high glucose (20 mmol/l)

concentration, Sirt1BKO cells exhibited reduced [Ca2+]i (Fig. 16c,d). To show that these changes

were not due to altered activity of the voltage-dependent calcium channels, direct

depolarisation was induced by 30 mmol/l KCl. This elicited the same level of calcium influx in

both Sirt1BKO and control cells (Fig. 16c,e). Thus, Sirt1 inactivation impairs glucose-stimulated

calcium influx without altering calcium-channel function.

47

Figure 16: Decreased ATP production and calcium influx in Sirt1-inactivated cells. (a) Glucose-stimulated ATP production in non-treated (blue bar) and 25µM Resveratrol-treated (green bar) MIN6 cells. N=2 per group. (b) Glucose-stimulated ATP production in non-treated (blue bar), 25µM Resveratrol-treated (green bar) and 25µM Sirtinol-treated (red bar) CD1 islets. n=2 per group. (c) Representative kinetic traces of the Fura2AM fluorescent signal from pooled Sirt1BKO (black trace) and control (grey trace) beta cells. (d, e) Summary of 20 mmol/l glucose- and KCl (30 mmol/l)-induced calcium uptakes in control (white bar) and Sirt1BKO (black bar) beta cells. n=3 independent experiments, and each experiment used ~30–100 cells from 3-4 mice per group. *p<0.05

48

5 Discussion

Beta cell-specific Sirt1 deletion causes impaired insulin secretion and glucose

intolerance in mice. In vitro, decreased GSIS from Sirt1BKO islets and SIRT1KD MIN6 cells is

associated with defective glucose sensing. By metabolic, molecular and gene analysis, we

uncovered a large panel of altered expression of mitochondria-related genes combined with

impaired glucose-induced MMP and decreased oxygen consumption. Our data show that SIRT1

is essential for mitochondrial function in pancreatic beta cells, which when dysfunctional, will

lead to downstream reduced calcium influx, the trigger for insulin release. In the absence of

SIRT1, beta cells can no longer perform the vital function of balancing fluctuations in glucose

uptake with insulin secretion. Therefore SIRT1’s actions in the beta cell appear to be

instrumental for metabolism.

While the importance of studying Sirt1 in a tissue-specific fashion is important to

understanding its effects on whole-body glucose homeostasis, there are compromises to

achieving this. The inducible Cre-Lox system is inherently subject to certain technical pitfalls

that must be considered. To begin, tamoxifen is the ligand used to bind the estrogen receptor

portion of the CreER fusion protein to induce translocation into the nucleus. In the context of

diabetes, this could pose a confounding effect as tamoxifen has been shown to act as a mild

uncoupler of mitochondrial oxidative phosphorylation[187] which could potentially affect

insulin secretion. Furthermore, tamoxifen has also been shown to reduce insulin sensitivity in

49

obese human patients[188]. Our lab has previously addressed tamoxifen’s effect on insulin

secretion from treated and non-treated islets which showed that the dose we used (125mg/kg)

does not yield a difference between the two groups. Similarly, glucose tolerance was assessed

in tamoxifen-injected and non-treated FloxSirt1 mice. It showed no difference in blood glucose

levels nor plasma insulin concentrations indicating that insulin resistance was not induced by

our tamoxifen treatment regimen.

Temporal control of deletion is predicated on sequestration of CreER fusion protein in

the cytosol. Estrogens are an endogenous ligand to the normal estrogen receptor, and would

therefore abolish any temporally-restricted gene deletion. In light of this, mutations were

incorporated into the ligand-binding domains of the CreER fusion protein such that activation

could only be achieved by synthetic but not natural estrogens[189, 190]. While this idea is

innovative, the imperfect nature of relatively new methodologies raises the question of CreER

“leakage” in this context. Liu et al. demonstrated this phenomenon by crossing RIP-CreER mice

to the loxP-Stop-lox-Rosa26-LacZ reporter strain, which revealed 42.9% lacZ staining in the

islets of 10-week-old untreated (tamoxifen) mice[191]. Although we do see a faint lower band

in our RT-PCR indicating that these tamoxifen-independent recombination events may be

occurring in our model, it is important to note that not all CreER strains are equal. In the same

study, Liu et al. also showed that the Pdx1-CreER mouse which was generated in the Melton lab

and used in our studies exhibit largely reduced leakiness[191]. Thus, Pdx1CreERxLoxSirt1 mice

closely approximate an overall acute pancreatic beta cell Sirt1 deletion.

50

Finally, the tissue-specific recombinase activity is dependent on the spatial exclusivity of

the promoter used to drive CreER expression. Recently, Wicksteed et al. examined the tissue-

distributional activity of the Pdx1 promoter by crossing Pdx1Cre mice to the Rosa26 report

strain. This group found that Cre activity in the hypothalamus, particularly orexin-expressing

neurons[180], could affect food intake, hepatic glucose production, and overall glucose and

energy homeostasis[192]. Whether these results can be generalized to all different strains of

Pdx1-Cre mice is still not known. Therefore it is incumbent upon the investigators using these

transgenic mice to investigate the level of Cre activity in the hypothalamus and other non-islet

beta cell tissues that could logically confound the phenotype of their models. We did address

this point and we showed by both RT-PCR and qPCR that Sirt1 levels were not affected in the

hypothalamus. We also showed this result in the intestine, which is another site of reported

Pdx1 activity[193-195], which could potentially affect the the release and/or functions of

incretin hormones. Thus, the phenotype of Pdx1CreERxLoxSirt1 mice is most probably due to

alterations in the islet beta cell and not other metabolic tissues.

Consistent with observations in whole-body Sirt1 knockout (Sirt1−/−) mice, we showed

that Sirt1BKO mice secrete significantly less insulin when challenged with glucose compared to

controls. Moreover, islets isolated from Sirt1−/− and Sirt1BKO mice both exhibit markedly

impaired insulin secretion when treated with stimulatory glucose concentration[78]. Due to the

heterogenous population of cells contained in an islet i.e. alpha and delta cells, Bordone et al.

and our group showed that in pure beta cell lines (MIN6 and INS1). siRNA-mediated SIRT1

knockdown significantly dampened insulin secretion thereby ruling out any possible paracrine

effect from non-beta cell neighbouring cells. Accordingly, beta cell overexpression (~18-fold) of

51

SIRT1 in BESTO mice enhanced insulin secretion [166]. Although this provides important proof-

of-principle, the dramatic Sirt1 overexpression may not offer the most physiologically relevant

scenario. This point of contention is addressed by the SirBACO mice who exhibit a much lower

level of Sirt1 overexpression (~twofold) and reap many metabolic benefits including resistance

against insulin desensitization and hyperglycemia. Of note, however, is that SirBACO mice do

not experience the same boosted insulin secretion which suggests that this effect is dose

dependent[168]. We used 25 µmol/l resveratrol to activate SIRT1 and subsequently boost

insulin secretion. This dosage was shown by Howitz et al. to enhance deactylase activity

~threefold[196], demonstrating that a relatively modest increase in SIRT1 activity is needed to

achieve increased GSIS. Our findings, along with previous reports, suggest a positive association

between SIRT1 and insulin secretion depending on the level of expression. It appears from

these observations that SIRT1’s actions on insulin resistance may apply to the whole organism

in an attempt to maintain glucose homeostasis before its effects on the beta cell set in. This

makes sense from a physiological point of view as it is metabolically favourable to maintain

insulin sensitivity as a means to lower blood glucose rather than hyper-secreting insulin. But

continuous over-feeding may eventually overwhelm SIRT1’s insulin sensitizing effects where in

the pancreatic beta cell, higher SIRT1 activity at this point will increase insulin secretion to

counteract the persisting hyperglycemia.

Sirt1−/− mice exhibit lower blood glucose ad libitum whereas Sirt1BKO mice do not [78].

At first glance, these results are counter-intuitive based on SIRT1’s protective role against

insulin resistance. However, these mice were not challenged with a high-fat diet, but rather,

regular chow. Morever, Sirt1-/- mice undergo Sirt1 recombination in the embryo which would

52

lower insulin secretion from birth resulting in potential compensation from peripheral tissues to

increase insulin sensitivity. This phenomenon of increased sensitization with concurrent

hypoinsulinemia has been observed in previous studies although the full mechanism is still yet

to be elucidated[197, 198]. Acute deletion in our Sirt1BKO mice does not allow enough time for

development of this adaption which could explain the lack of any difference in plasma insulin

levels ad libitum compared to controls. In addition, Sirt1BKO mice and controls were

maintained on a regular chow diet, not high fat, which does not act to stress the mice enough

to reveal the expected lower levels of plasma insulin in Sirt1BKOs ad libitum. Intriguingly, SIRT1

gain-of-function mice do not present with increased insulin sensitivity unless stressed with a

high-fat diet[168], suggesting that SIRT1 counteracts rises in insulin resistance but not

necessarily lowering resistance from a basal level.

It has been shown by others that SIRT1 acts to halt beta cell expansion by deacetylation

of FOXO1, which acts as a molecular handbrake on proliferation [150]. However, our data

shows no difference in beta cell area, which suggests that this pathway does not cause

decreased GSIS, at least in our acute Sirt1 inactivation model. Furthermore, the islet and beta

cell area of BESTO mice did not show any differences compared to controls. SIRT1’s prevention

of beta cell proliferation may reinforce the idea that the organism attempts to first maintain

peripheral insulin sensitivity before exhausting beta cell function. There is an arbitrarily

classified phase in the progression towards T2D, stage 2, whereby hyperglycemia can vary while

the beta cell is adapting without any changes to beta cell mass[162]. This may correlate with

the stage in which Sirt1 is promoting insulin secretion by increasing beta cell function without

increasing mass expansion. Our studies are relatively acute 5-10 days after Sirt1 deletion, which

53

may be too short of a time span to observe any changes to beta cell mass. Alternatively, the

findings by Bastien-Dionne et al. could simply be due to an artefact of the supraphysiological

concentrations (10nmol/l) used in their studies compared to those concentrations in the blood

typically after a meal (approximately 50pmol/l)[199]. The role of Sirt1 in regulating beta cell

mass is still in its infancy and more work will need to be done to understand what factors are

involved and how these events change over time.

Previous studies have linked Sirt1 knockdown with increased expression of Ucp2, which

is thought to cause increased proton leakage and hence decreased insulin secretion [78, 166].

Interestingly, while we show impaired GSIS, this was not associated with increased levels of

Ucp2 mRNA. While mRNA levels may not reflect protein levels accurately, we did measure

proton leakage in our oxygen consumption rate experiments which showed no difference in the

level of oligomycin-treated OCR. This suggested that UCP2 activity was not affected by SIRT1

knockdown in our SIRT1KD MIN6 model. We did, however, observe altered expression of a

large number of other mitochondria-related genes. Mitochondria must efficiently harvest

carbon derivatives of glucose to generate a proton gradient that drives the production of ATP

[200]. Without properly functioning mitochondria, the beta cell loses its exquisite ability to fine-

tune glucose uptake to insulin release. In cells lacking mitochondrial ATP production, GSIS is

abolished [201, 202]. In Sirt1-inactivated cells, reduced MMP and OCR are reflective of impaired

mitochondrial function, which likely results in reduced ATP production. We did not explicitly

examine the level of ATP in our Sirt1BKO islets and SIRT1KD MIN6 cells however, we’ve

localized the defect to mitochondrial respiration which we are confident will not only affect ATP

levels in our model but the impairments would manifest downstream leading to reduced insulin

54

secretion. Importantly, direct depolarisation by KCl could restore insulin secretion to control

levels [202] indicating that depolarization and calcium channel activity is intact in Sirt1

inactivated models and rendering it highly likely that ATP levels are decreased. These findings

demonstrate that ATP from mitochondria was crucial for depolarisation to induce Ca2+ influx

through voltage-dependent calcium channels. Reduced [Ca2+]i

A study in rat insulinoma beta cells (INS-1E) showed that enhanced SIRT1 activity

upregulated Glut2, Gck, Pdx1 and Tfam, a short list of genes that could affect beta cell function

without necessarily altering proton leak. Rather, these changes enhanced glycolytic flux, which

produced more mitochondrial substrates and ultimately boosted insulin secretion[203]. As

expected, the opposite is true: decreased SIRT1 activity resulted in reduced insulin secretion.

However, the altered genes in our SIRT1KD cells were not the same as those in INS-1E. Our

findings highlight novel genes under SIRT1 regulation in the beta cell. Interestingly, in SIRT1KD

cells, the mitochondrial master regulator, Pgc-1a, was upregulated more than twofold at the

mRNA level compared with controls. At first glance, these findings are counter-intuitive, as

increased Pgc-1a should boost mitochondrial function and biogenesis. Indeed, a number of

studies have shown PGC-1α caused increased mitochondrial respiration and production[204-

206], which in the beta cell would presumably result in enhanced insulin secretion. We can

offer three possibilities to explain this; first, increased Pgc-1a expression is a compensatory

mechanism by which levels of mitochondrial function are sustained when SIRT1 is inactive.

in Sirt1BKO cells in response to

high glucose levels supports this notion. Because Ucp2 expression could not adequately

account for the changes we observed, we were prompted to search for other genes under

SIRT1’s control that could.

55

Second, PGC-1α’s transcription and translation are increased to compensate in response to

impaired mitochondrial bioenergetics, however, in the absence of SIRT1, PGC-1a is left

acetylated and inactivate. Remarkably, SIRT1KD cells showed increased levels of acetylated

lysine in whole cell lysates. Lee and colleagues observed that SIRT1 protected pancreatic beta

cells from cytokine-induced damage by deacetylation-dependent repression of nuclear factor-

κB [155] which corroborates the idea that loss of SIRT1-mediated substrate deacetylation is

associated with beta cell dysfunction. Finally increased Pgc-1a actually contributes to beta-cell

dysfunction, distinct from its roles in other tissues. In fact, Yoon et al. showed that elevated

expression of PGC-1α in islets caused a marked decrease in insulin secretion by suppressing key

genes involved in glucose metabolism [207]. In any event we can infer that these alterations in

Sirt1 and Pgc-1a cause a disruption of mitochondrial dynamics, which affects their respiration

efficiency.

Age is one of the single largest risk factors for the development of type 2 diabetes[208].

The pathogenesis of type 2 diabetes has been linked to an age-related decline in pancreatic

levels of NAD+, the cofactor required for SIRT1 deacetylation [209]. This would theoretically

lead to declining SIRT1 activity mirroring the effects of Sirt1 inactivation. It stands to reason

then that if SIRT1 is orchestrating mitochondrial processes, then mitochondrial function should

decline with age as well. These observations are similar to those made in diabetic patients who

harboured mitochondrial DNA mutations and presented with pronounced age-dependent

deterioration of beta cell function [210]. In rodent models, these mitochondrial deficits could

be improved, however, by administering resveratrol, a small-molecule polyphenol which

requires SIRT1 for its action[182, 203]. In doing so, Resveratrol improved metabolic indices in

56

mice, curtailing the progression of type 2 diabetes [211, 212]. We showed that in our Sirt1BKO

islets that Resveratrol had no effect in enhancing insulin secretion whereas this effect was

observed in wildtype islets containing intact SIRT1. Very recently, Hubbard et al. provided

evidence for the mechanism of Resveratrol’s and other sirtuin-activating compounds (STACs) on

SIRT1 activity. They showed that a conserved amino acid in the N-terminal of SIRT1, Glu230, was

required for activation by all STACs and that a glutamine-to-lysine mutation in this position

significantly dampened these enhanced effects[213]. This has important implications for the

general population, especially those individuals who are living with diabetes or are currently in

the pre-diabetes stage. Any mutation or genetic polymorphism possessed by an individual that

changes the conserved Glu230

amino acid may not reap the benefits from any therapeutic

and/or nutritional intervention targeting sirtuins.

57

6 Conclusion

Our study attempts to elucidate the molecular basis of SIRT1’s role in insulin secretion. We

demonstrate that SIRT1 regulates a complex network of mitochondria-related genes, whose

levels are disrupted by Sirt1 deletion. This impaired mitochondrial function ultimately had a

negative impact on GSIS. At the apex of type 2 diabetes is the inability to secrete insulin

adequately but, where this mechanism fails, SIRT1 offers a new avenue of therapeutic

possibilities.

58

7 References

1. Danaei, G., et al., National, regional, and global trends in systolic blood pressure since 1980: systematic analysis of health examination surveys and epidemiological studies with 786 country-years and 5.4 million participants. Lancet, 2011. 377(9765): p. 568-77.

2. Lin, Y. and Z. Sun, Current views on type 2 diabetes. J Endocrinol, 2010. 204(1): p. 1-11. 3. Zimmet, P., K.G. Alberti, and J. Shaw, Global and societal implications of the diabetes

epidemic. Nature, 2001. 414(6865): p. 782-7. 4. Harris, M.I., et al., Prevalence of diabetes and impaired glucose tolerance and plasma

glucose levels in U.S. population aged 20-74 yr. Diabetes, 1987. 36(4): p. 523-34. 5. Ferrannini, E., et al., Mode of onset of type 2 diabetes from normal or impaired glucose

tolerance. Diabetes, 2004. 53(1): p. 160-5. 6. Nichols, G.A., T.A. Hillier, and J.B. Brown, Progression from newly acquired impaired

fasting glusose to type 2 diabetes. Diabetes Care, 2007. 30(2): p. 228-33. 7. Venables, M.C. and A.E. Jeukendrup, Physical inactivity and obesity: links with insulin

resistance and type 2 diabetes mellitus. Diabetes Metab Res Rev, 2009. 25 Suppl 1: p. S18-23.

8. Polonsky, K.S., et al., Quantitative study of insulin secretion and clearance in normal and obese subjects. J Clin Invest, 1988. 81(2): p. 435-41.

9. Arslanian, S.A., et al., Hyperinsulinemia in african-american children: decreased insulin clearance and increased insulin secretion and its relationship to insulin sensitivity. Diabetes, 2002. 51(10): p. 3014-9.

10. Weyer, C., et al., Insulin resistance and insulin secretory dysfunction are independent predictors of worsening of glucose tolerance during each stage of type 2 diabetes development. Diabetes Care, 2001. 24(1): p. 89-94.

11. Weir, G.C., et al., Beta-cell adaptation and decompensation during the progression of diabetes. Diabetes, 2001. 50 Suppl 1: p. S154-9.

12. Koulajian, K., et al., NADPH oxidase inhibition prevents beta cell dysfunction induced by prolonged elevation of oleate in rodents. Diabetologia, 2013. 56(5): p. 1078-87.

13. Butler, A.E., et al., Beta-cell deficit and increased beta-cell apoptosis in humans with type 2 diabetes. Diabetes, 2003. 52(1): p. 102-10.

14. Matschinsky, F.M., B. Glaser, and M.A. Magnuson, Pancreatic beta-cell glucokinase: closing the gap between theoretical concepts and experimental realities. Diabetes, 1998. 47(3): p. 307-15.

15. Rorsman, P. and M. Braun, Regulation of insulin secretion in human pancreatic islets. Annu Rev Physiol, 2013. 75: p. 155-79.

16. Matschinsky, F.M., Banting Lecture 1995. A lesson in metabolic regulation inspired by the glucokinase glucose sensor paradigm. Diabetes, 1996. 45(2): p. 223-41.

59

17. Stoffel, M., et al., Human glucokinase gene: isolation, characterization, and identification of two missense mutations linked to early-onset non-insulin-dependent (type 2) diabetes mellitus. Proc Natl Acad Sci U S A, 1992. 89(16): p. 7698-702.

18. Akram, M., Mini-review on Glycolysis and Cancer. J Cancer Educ, 2013. 19. Fernie, A.R., F. Carrari, and L.J. Sweetlove, Respiratory metabolism: glycolysis, the TCA

cycle and mitochondrial electron transport. Curr Opin Plant Biol, 2004. 7(3): p. 254-61. 20. Papa, S., et al., The oxidative phosphorylation system in mammalian mitochondria. Adv

Exp Med Biol, 2012. 942: p. 3-37. 21. Schagger, H. and K. Pfeiffer, Supercomplexes in the respiratory chains of yeast and

mammalian mitochondria. EMBO J, 2000. 19(8): p. 1777-83. 22. Mitchell, P., Coupling of phosphorylation to electron and hydrogen transfer by a chemi-

osmotic type of mechanism. Nature, 1961. 191: p. 144-8. 23. Papa, S., Proton translocation reactions in the respiratory chains. Biochim Biophys Acta,

1976. 456(1): p. 39-84. 24. Sapra, R., K. Bagramyan, and M.W. Adams, A simple energy-conserving system: proton

reduction coupled to proton translocation. Proc Natl Acad Sci U S A, 2003. 100(13): p. 7545-50.

25. Porcelli, D., et al., The nuclear OXPHOS genes in insecta: a common evolutionary origin, a common cis-regulatory motif, a common destiny for gene duplicates. BMC Evol Biol, 2007. 7: p. 215.

26. Papa, S., et al., Mammalian complex I: a regulable and vulnerable pacemaker in mitochondrial respiratory function. Biochim Biophys Acta, 2008. 1777(7-8): p. 719-28.

27. Hirst, J., et al., The nuclear encoded subunits of complex I from bovine heart mitochondria. Biochim Biophys Acta, 2003. 1604(3): p. 135-50.

28. Cocco, T., et al., Control of OXPHOS efficiency by complex I in brain mitochondria. Neurobiol Aging, 2009. 30(4): p. 622-9.

29. Schagger, H., et al., Ubiquinol-cytochrome-c reductase from human and bovine mitochondria. Methods Enzymol, 1995. 260: p. 82-96.

30. Matsuno-Yagi, A. and Y. Hatefi, Ubiquinol:cytochrome c oxidoreductase (complex III). Effect of inhibitors on cytochrome b reduction in submitochondrial particles and the role of ubiquinone in complex III. J Biol Chem, 2001. 276(22): p. 19006-11.

31. Wikstrom, M., Mechanism of proton translocation by cytochrome c oxidase: a new four-stroke histidine cycle. Biochim Biophys Acta, 2000. 1458(1): p. 188-98.

32. Brzezinski, P. and R.B. Gennis, Cytochrome c oxidase: exciting progress and remaining mysteries. J Bioenerg Biomembr, 2008. 40(5): p. 521-31.

33. Tsukihara, T., et al., The low-spin heme of cytochrome c oxidase as the driving element of the proton-pumping process. Proc Natl Acad Sci U S A, 2003. 100(26): p. 15304-9.

34. Yoshikawa, S., et al., Proton pumping mechanism of bovine heart cytochrome c oxidase. Biochim Biophys Acta, 2006. 1757(9-10): p. 1110-6.

35. Kabaleeswaran, V., et al., Novel features of the rotary catalytic mechanism revealed in the structure of yeast F1 ATPase. EMBO J, 2006. 25(22): p. 5433-42.

36. Boyer, P.D., The ATP synthase--a splendid molecular machine. Annu Rev Biochem, 1997. 66: p. 717-49.

60

37. Deckers-Hebestreit, G. and K. Altendorf, The F0F1-type ATP synthases of bacteria: structure and function of the F0 complex. Annu Rev Microbiol, 1996. 50: p. 791-824.

38. de Weille, J., et al., ATP-sensitive K+ channels that are blocked by hypoglycemia-inducing sulfonylureas in insulin-secreting cells are activated by galanin, a hyperglycemia-inducing hormone. Proc Natl Acad Sci U S A, 1988. 85(4): p. 1312-6.

39. Misler, S., et al., Metabolite-regulated ATP-sensitive K+ channel in human pancreatic islet cells. Diabetes, 1989. 38(4): p. 422-7.

40. Misler, S., et al., Electrophysiology of stimulus-secretion coupling in human beta-cells. Diabetes, 1992. 41(10): p. 1221-8.

41. Tarasov, A., J. Dusonchet, and F. Ashcroft, Metabolic regulation of the pancreatic beta-cell ATP-sensitive K+ channel: a pas de deux. Diabetes, 2004. 53 Suppl 3: p. S113-22.

42. Braun, M., et al., Voltage-gated ion channels in human pancreatic beta-cells: electrophysiological characterization and role in insulin secretion. Diabetes, 2008. 57(6): p. 1618-28.

43. Yang, S.N. and P.O. Berggren, The role of voltage-gated calcium channels in pancreatic beta-cell physiology and pathophysiology. Endocr Rev, 2006. 27(6): p. 621-76.

44. Henquin, J.C., et al., Hierarchy of the beta-cell signals controlling insulin secretion. Eur J Clin Invest, 2003. 33(9): p. 742-50.

45. Newgard, C.B. and J.D. McGarry, Metabolic coupling factors in pancreatic beta-cell signal transduction. Annu Rev Biochem, 1995. 64: p. 689-719.

46. Petersen, K.F., et al., Mitochondrial dysfunction in the elderly: possible role in insulin resistance. Science, 2003. 300(5622): p. 1140-2.

47. Kim, J.A., Y. Wei, and J.R. Sowers, Role of mitochondrial dysfunction in insulin resistance. Circ Res, 2008. 102(4): p. 401-14.

48. Patti, M.E. and S. Corvera, The role of mitochondria in the pathogenesis of type 2 diabetes. Endocr Rev, 2010. 31(3): p. 364-95.

49. Galgani, J.E., C. Moro, and E. Ravussin, Metabolic flexibility and insulin resistance. Am J Physiol Endocrinol Metab, 2008. 295(5): p. E1009-17.

50. DeFronzo, R.A. and D. Tripathy, Skeletal muscle insulin resistance is the primary defect in type 2 diabetes. Diabetes Care, 2009. 32 Suppl 2: p. S157-63.

51. Simoneau, J.A., et al., Skeletal muscle glycolytic and oxidative enzyme capacities are determinants of insulin sensitivity and muscle composition in obese women. FASEB J, 1995. 9(2): p. 273-8.

52. Savage, D.B., K.F. Petersen, and G.I. Shulman, Disordered lipid metabolism and the pathogenesis of insulin resistance. Physiol Rev, 2007. 87(2): p. 507-20.

53. Koves, T.R., et al., Mitochondrial overload and incomplete fatty acid oxidation contribute to skeletal muscle insulin resistance. Cell Metab, 2008. 7(1): p. 45-56.

54. Houstis, N., E.D. Rosen, and E.S. Lander, Reactive oxygen species have a causal role in multiple forms of insulin resistance. Nature, 2006. 440(7086): p. 944-8.

55. Ritov, V.B., et al., Deficiency of subsarcolemmal mitochondria in obesity and type 2 diabetes. Diabetes, 2005. 54(1): p. 8-14.

56. Hoeks, J. and P. Schrauwen, Muscle mitochondria and insulin resistance: a human perspective. Trends Endocrinol Metab, 2012. 23(9): p. 444-50.

61

57. Kelley, D.E., et al., Dysfunction of mitochondria in human skeletal muscle in type 2 diabetes. Diabetes, 2002. 51(10): p. 2944-50.

58. Mogensen, M., et al., Mitochondrial respiration is decreased in skeletal muscle of patients with type 2 diabetes. Diabetes, 2007. 56(6): p. 1592-9.

59. Szendroedi, J., et al., Impaired mitochondrial function and insulin resistance of skeletal muscle in mitochondrial diabetes. Diabetes Care, 2009. 32(4): p. 677-9.

60. Rabol, R., et al., Effect of hyperglycemia on mitochondrial respiration in type 2 diabetes. J Clin Endocrinol Metab, 2009. 94(4): p. 1372-8.

61. Mootha, V.K., et al., PGC-1alpha-responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat Genet, 2003. 34(3): p. 267-73.

62. Crunkhorn, S., et al., Peroxisome proliferator activator receptor gamma coactivator-1 expression is reduced in obesity: potential pathogenic role of saturated fatty acids and p38 mitogen-activated protein kinase activation. J Biol Chem, 2007. 282(21): p. 15439-50.

63. Sparks, L.M., et al., A high-fat diet coordinately downregulates genes required for mitochondrial oxidative phosphorylation in skeletal muscle. Diabetes, 2005. 54(7): p. 1926-33.

64. van Tienen, F.H., et al., Physical activity is the key determinant of skeletal muscle mitochondrial function in type 2 diabetes. J Clin Endocrinol Metab, 2012. 97(9): p. 3261-9.

65. Cannon, B. and J. Nedergaard, Brown adipose tissue: function and physiological significance. Physiol Rev, 2004. 84(1): p. 277-359.

66. Porter, C., E. Borsheim, and L.S. Sidossis, Does adipose tissue thermogenesis play a role in metabolic health? J Obes, 2013. 2013: p. 204094.

67. van Marken Lichtenbelt, W.D., et al., Cold-activated brown adipose tissue in healthy men. N Engl J Med, 2009. 360(15): p. 1500-8.

68. Valerio, A., et al., TNF-alpha downregulates eNOS expression and mitochondrial biogenesis in fat and muscle of obese rodents. J Clin Invest, 2006. 116(10): p. 2791-8.

69. Kusminski, C.M. and P.E. Scherer, Mitochondrial dysfunction in white adipose tissue. Trends Endocrinol Metab, 2012. 23(9): p. 435-43.

70. Choo, H.J., et al., Mitochondria are impaired in the adipocytes of type 2 diabetic mice. Diabetologia, 2006. 49(4): p. 784-91.

71. Hasan, N.M., et al., Impaired anaplerosis and insulin secretion in insulinoma cells caused by small interfering RNA-mediated suppression of pyruvate carboxylase. J Biol Chem, 2008. 283(42): p. 28048-59.

72. Leloup, C., et al., Mitochondrial reactive oxygen species are obligatory signals for glucose-induced insulin secretion. Diabetes, 2009. 58(3): p. 673-81.

73. Silva, J.P., et al., Impaired insulin secretion and beta-cell loss in tissue-specific knockout mice with mitochondrial diabetes. Nat Genet, 2000. 26(3): p. 336-40.

74. Hou, L., et al., Pseudolaric acid B inhibits inducible cyclooxygenase-2 expression via downregulation of the NF-kappaB pathway in HT-29 cells. J Cancer Res Clin Oncol, 2012.

75. Lin, N., et al., Mitochondrial reactive oxygen species (ROS) inhibition ameliorates palmitate-induced INS-1 beta cell death. Endocrine, 2012. 42(1): p. 107-17.

62

76. Produit-Zengaffinen, N., et al., Increasing uncoupling protein-2 in pancreatic beta cells does not alter glucose-induced insulin secretion but decreases production of reactive oxygen species. Diabetologia, 2007. 50(1): p. 84-93.

77. Zhang, C.Y., et al., Uncoupling protein-2 negatively regulates insulin secretion and is a major link between obesity, beta cell dysfunction, and type 2 diabetes. Cell, 2001. 105(6): p. 745-55.

78. Bordone, L., et al., Sirt1 regulates insulin secretion by repressing UCP2 in pancreatic beta cells. PLoS Biol, 2006. 4(2): p. e31.

79. Braunstein, M., et al., Transcriptional silencing in yeast is associated with reduced nucleosome acetylation. Genes Dev, 1993. 7(4): p. 592-604.

80. Thompson, J.S., X. Ling, and M. Grunstein, Histone H3 amino terminus is required for telomeric and silent mating locus repression in yeast. Nature, 1994. 369(6477): p. 245-7.

81. Kaeberlein, M., M. McVey, and L. Guarente, The SIR2/3/4 complex and SIR2 alone promote longevity in Saccharomyces cerevisiae by two different mechanisms. Genes Dev, 1999. 13(19): p. 2570-80.

82. Garcia-Salcedo, J.A., et al., A chromosomal SIR2 homologue with both histone NAD-dependent ADP-ribosyltransferase and deacetylase activities is involved in DNA repair in Trypanosoma brucei. EMBO J, 2003. 22(21): p. 5851-62.

83. Kruszewski, M. and I. Szumiel, Sirtuins (histone deacetylases III) in the cellular response to DNA damage--facts and hypotheses. DNA Repair (Amst), 2005. 4(11): p. 1306-13.

84. Gallo, C.M., D.L. Smith, Jr., and J.S. Smith, Nicotinamide clearance by Pnc1 directly regulates Sir2-mediated silencing and longevity. Mol Cell Biol, 2004. 24(3): p. 1301-12.

85. Lin, S.J., P.A. Defossez, and L. Guarente, Requirement of NAD and SIR2 for life-span extension by calorie restriction in Saccharomyces cerevisiae. Science, 2000. 289(5487): p. 2126-8.

86. Imai, S., et al., Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase. Nature, 2000. 403(6771): p. 795-800.

87. Lin, S.J., et al., Calorie restriction extends Saccharomyces cerevisiae lifespan by increasing respiration. Nature, 2002. 418(6895): p. 344-8.

88. Brachmann, C.B., et al., The SIR2 gene family, conserved from bacteria to humans, functions in silencing, cell cycle progression, and chromosome stability. Genes Dev, 1995. 9(23): p. 2888-902.

89. Frye, R.A., Characterization of five human cDNAs with homology to the yeast SIR2 gene: Sir2-like proteins (sirtuins) metabolize NAD and may have protein ADP-ribosyltransferase activity. Biochem Biophys Res Commun, 1999. 260(1): p. 273-9.

90. Frye, R.A., Phylogenetic classification of prokaryotic and eukaryotic Sir2-like proteins. Biochem Biophys Res Commun, 2000. 273(2): p. 793-8.

91. Stunkel, W. and R.M. Campbell, Sirtuin 1 (SIRT1): the misunderstood HDAC. J Biomol Screen, 2011. 16(10): p. 1153-69.

92. Michishita, E., et al., Evolutionarily conserved and nonconserved cellular localizations and functions of human SIRT proteins. Mol Biol Cell, 2005. 16(10): p. 4623-35.

93. Haigis, M.C. and D.A. Sinclair, Mammalian sirtuins: biological insights and disease relevance. Annu Rev Pathol, 2010. 5: p. 253-95.

63

94. Baur, J.A., et al., Are sirtuins viable targets for improving healthspan and lifespan? Nat Rev Drug Discov, 2012. 11(6): p. 443-61.

95. Liou, G.G., et al., Assembly of the SIR complex and its regulation by O-acetyl-ADP-ribose, a product of NAD-dependent histone deacetylation. Cell, 2005. 121(4): p. 515-27.

96. Vaquero, A., et al., Human SirT1 interacts with histone H1 and promotes formation of facultative heterochromatin. Mol Cell, 2004. 16(1): p. 93-105.

97. Avalos, J.L., et al., Structure of a Sir2 enzyme bound to an acetylated p53 peptide. Mol Cell, 2002. 10(3): p. 523-35.

98. Blander, G. and L. Guarente, The Sir2 family of protein deacetylases. Annu Rev Biochem, 2004. 73: p. 417-35.

99. Yamamoto, H., K. Schoonjans, and J. Auwerx, Sirtuin functions in health and disease. Mol Endocrinol, 2007. 21(8): p. 1745-55.

100. Li, X. and N. Kazgan, Mammalian sirtuins and energy metabolism. Int J Biol Sci, 2011. 7(5): p. 575-87.

101. Yu, J. and J. Auwerx, The role of sirtuins in the control of metabolic homeostasis. Ann N Y Acad Sci, 2009. 1173 Suppl 1: p. E10-9.

102. Nogueiras, R., et al., Sirtuin 1 and sirtuin 3: physiological modulators of metabolism. Physiol Rev, 2012. 92(3): p. 1479-514.

103. Cheng, H.L., et al., Developmental defects and p53 hyperacetylation in Sir2 homolog (SIRT1)-deficient mice. Proc Natl Acad Sci U S A, 2003. 100(19): p. 10794-9.

104. Vaziri, H., et al., hSIR2(SIRT1) functions as an NAD-dependent p53 deacetylase. Cell, 2001. 107(2): p. 149-59.

105. Zilfou, J.T. and S.W. Lowe, Tumor suppressive functions of p53. Cold Spring Harb Perspect Biol, 2009. 1(5): p. a001883.

106. MacFarlane, M. and A.C. Williams, Apoptosis and disease: a life or death decision. EMBO Rep, 2004. 5(7): p. 674-8.

107. Jeong, J., et al., SIRT1 promotes DNA repair activity and deacetylation of Ku70. Exp Mol Med, 2007. 39(1): p. 8-13.

108. Yuan, Z., et al., SIRT1 regulates the function of the Nijmegen breakage syndrome protein. Mol Cell, 2007. 27(1): p. 149-62.

109. Li, K., et al., Regulation of WRN protein cellular localization and enzymatic activities by SIRT1-mediated deacetylation. J Biol Chem, 2008. 283(12): p. 7590-8.

110. Ming, M., et al., Regulation of global genome nucleotide excision repair by SIRT1 through xeroderma pigmentosum C. Proc Natl Acad Sci U S A, 2010. 107(52): p. 22623-8.

111. Wang, R.H., et al., Interplay among BRCA1, SIRT1, and Survivin during BRCA1-associated tumorigenesis. Mol Cell, 2008. 32(1): p. 11-20.

112. Benayoun, B.A., et al., Transcription factor FOXL2 protects granulosa cells from stress and delays cell cycle: role of its regulation by the SIRT1 deacetylase. Hum Mol Genet, 2011. 20(9): p. 1673-86.

113. Giannakou, M.E. and L. Partridge, The interaction between FOXO and SIRT1: tipping the balance towards survival. Trends Cell Biol, 2004. 14(8): p. 408-12.

114. Liu, Y., et al., A fasting inducible switch modulates gluconeogenesis via activator/coactivator exchange. Nature, 2008. 456(7219): p. 269-73.

64

115. Sasaki, T., et al., Induction of hypothalamic Sirt1 leads to cessation of feeding via agouti-related peptide. Endocrinology, 2010. 151(6): p. 2556-66.

116. Picard, F., et al., Sirt1 promotes fat mobilization in white adipocytes by repressing PPAR-gamma. Nature, 2004. 429(6993): p. 771-6.

117. Hou, X., et al., SIRT1 regulates hepatocyte lipid metabolism through activating AMP-activated protein kinase. J Biol Chem, 2008. 283(29): p. 20015-26.

118. Toledo, F.G., A.D. Sniderman, and D.E. Kelley, Influence of hepatic steatosis (fatty liver) on severity and composition of dyslipidemia in type 2 diabetes. Diabetes Care, 2006. 29(8): p. 1845-50.

119. Purushotham, A., et al., Hepatocyte-specific deletion of SIRT1 alters fatty acid metabolism and results in hepatic steatosis and inflammation. Cell Metab, 2009. 9(4): p. 327-38.

120. Li, X., et al., SIRT1 deacetylates and positively regulates the nuclear receptor LXR. Mol Cell, 2007. 28(1): p. 91-106.

121. Defour, A., et al., Sirtuin 1 regulates SREBP-1c expression in a LXR-dependent manner in skeletal muscle. PLoS One, 2012. 7(9): p. e43490.

122. Walker, A.K., et al., Conserved role of SIRT1 orthologs in fasting-dependent inhibition of the lipid/cholesterol regulator SREBP. Genes Dev, 2010. 24(13): p. 1403-17.

123. Rodgers, J.T., et al., Nutrient control of glucose homeostasis through a complex of PGC-1alpha and SIRT1. Nature, 2005. 434(7029): p. 113-8.

124. Rodgers, J.T. and P. Puigserver, Fasting-dependent glucose and lipid metabolic response through hepatic sirtuin 1. Proc Natl Acad Sci U S A, 2007. 104(31): p. 12861-6.

125. Shoelson, S.E., L. Herrero, and A. Naaz, Obesity, inflammation, and insulin resistance. Gastroenterology, 2007. 132(6): p. 2169-80.

126. Sun, C., et al., SIRT1 improves insulin sensitivity under insulin-resistant conditions by repressing PTP1B. Cell Metab, 2007. 6(4): p. 307-19.

127. Zhang, J., The direct involvement of SirT1 in insulin-induced insulin receptor substrate-2 tyrosine phosphorylation. J Biol Chem, 2007. 282(47): p. 34356-64.

128. Wang, R.H., et al., Hepatic Sirt1 deficiency in mice impairs mTorc2/Akt signaling and results in hyperglycemia, oxidative damage, and insulin resistance. J Clin Invest, 2011. 121(11): p. 4477-90.

129. Yamauchi, T., et al., The fat-derived hormone adiponectin reverses insulin resistance associated with both lipoatrophy and obesity. Nat Med, 2001. 7(8): p. 941-6.

130. Qiao, L. and J. Shao, SIRT1 regulates adiponectin gene expression through Foxo1-C/enhancer-binding protein alpha transcriptional complex. J Biol Chem, 2006. 281(52): p. 39915-24.

131. Zhu, M., et al., Circulating adiponectin levels increase in rats on caloric restriction: the potential for insulin sensitization. Exp Gerontol, 2004. 39(7): p. 1049-59.

132. Smith, J.J., et al., Small molecule activators of SIRT1 replicate signaling pathways triggered by calorie restriction in vivo. BMC Syst Biol, 2009. 3: p. 31.

133. Cohen, H.Y., et al., Calorie restriction promotes mammalian cell survival by inducing the SIRT1 deacetylase. Science, 2004. 305(5682): p. 390-2.

134. Cai, D., et al., Local and systemic insulin resistance resulting from hepatic activation of IKK-beta and NF-kappaB. Nat Med, 2005. 11(2): p. 183-90.

65

135. Yuan, M., et al., Reversal of obesity- and diet-induced insulin resistance with salicylates or targeted disruption of Ikkbeta. Science, 2001. 293(5535): p. 1673-7.

136. Shulman, J.M. and J.A. Schneider, Molecular mechanisms of cortical degeneration in Parkinson disease. Neurology, 2012. 79(17): p. 1750-1.

137. Hotamisligil, G.S., et al., IRS-1-mediated inhibition of insulin receptor tyrosine kinase activity in TNF-alpha- and obesity-induced insulin resistance. Science, 1996. 271(5249): p. 665-8.

138. Yoshizaki, T., et al., SIRT1 exerts anti-inflammatory effects and improves insulin sensitivity in adipocytes. Mol Cell Biol, 2009. 29(5): p. 1363-74.

139. Yeung, F., et al., Modulation of NF-kappaB-dependent transcription and cell survival by the SIRT1 deacetylase. EMBO J, 2004. 23(12): p. 2369-80.

140. Baldwin, A.S., Jr., The NF-kappa B and I kappa B proteins: new discoveries and insights. Annu Rev Immunol, 1996. 14: p. 649-83.

141. Tissenbaum, H.A. and L. Guarente, Increased dosage of a sir-2 gene extends lifespan in Caenorhabditis elegans. Nature, 2001. 410(6825): p. 227-30.

142. Daitoku, H., et al., Silent information regulator 2 potentiates Foxo1-mediated transcription through its deacetylase activity. Proc Natl Acad Sci U S A, 2004. 101(27): p. 10042-7.

143. Nakae, J., et al., The forkhead transcription factor Foxo1 (Fkhr) confers insulin sensitivity onto glucose-6-phosphatase expression. J Clin Invest, 2001. 108(9): p. 1359-67.

144. Farmer, S.R., The forkhead transcription factor Foxo1: a possible link between obesity and insulin resistance. Mol Cell, 2003. 11(1): p. 6-8.

145. Kitamura, T., et al., The forkhead transcription factor Foxo1 links insulin signaling to Pdx1 regulation of pancreatic beta cell growth. J Clin Invest, 2002. 110(12): p. 1839-47.

146. Kitamura, Y.I., et al., FoxO1 protects against pancreatic beta cell failure through NeuroD and MafA induction. Cell Metab, 2005. 2(3): p. 153-63.

147. Gu, C., et al., Pancreatic beta cells require NeuroD to achieve and maintain functional maturity. Cell Metab, 2010. 11(4): p. 298-310.

148. Huang, H.P., et al., Neogenesis of beta-cells in adult BETA2/NeuroD-deficient mice. Mol Endocrinol, 2002. 16(3): p. 541-51.

149. Kaneto, H., et al., Role of MafA in pancreatic beta-cells. Adv Drug Deliv Rev, 2009. 61(7-8): p. 489-96.

150. Bastien-Dionne, P.O., et al., Glucagon-like peptide 1 inhibits the sirtuin deacetylase SirT1 to stimulate pancreatic beta-cell mass expansion. Diabetes, 2011. 60(12): p. 3217-22.

151. Eizirik, D.L., M.L. Colli, and F. Ortis, The role of inflammation in insulitis and beta-cell loss in type 1 diabetes. Nat Rev Endocrinol, 2009. 5(4): p. 219-26.

152. Donath, M.Y., et al., Islet inflammation in type 2 diabetes: from metabolic stress to therapy. Diabetes Care, 2008. 31 Suppl 2: p. S161-4.

153. Collier, J.J., et al., Pancreatic beta-cell death in response to pro-inflammatory cytokines is distinct from genuine apoptosis. PLoS One, 2011. 6(7): p. e22485.

154. Li, Q. and I.M. Verma, NF-kappaB regulation in the immune system. Nat Rev Immunol, 2002. 2(10): p. 725-34.

66

155. Lee, J.H., et al., Overexpression of SIRT1 protects pancreatic beta-cells against cytokine toxicity by suppressing the nuclear factor-kappaB signaling pathway. Diabetes, 2009. 58(2): p. 344-51.

156. Szkudelski, T., The mechanism of alloxan and streptozotocin action in B cells of the rat pancreas. Physiol Res, 2001. 50(6): p. 537-46.

157. Wu, J., et al., WldS enhances insulin transcription and secretion via a SIRT1-dependent pathway and improves glucose homeostasis. Diabetes, 2011. 60(12): p. 3197-207.

158. Biason-Lauber, A., et al., Identification of a SIRT1 mutation in a family with type 1 diabetes. Cell Metab, 2013. 17(3): p. 448-55.

159. van Belle, T.L., K.T. Coppieters, and M.G. von Herrath, Type 1 diabetes: etiology, immunology, and therapeutic strategies. Physiol Rev, 2011. 91(1): p. 79-118.

160. Bassuny, W.M., et al., A functional polymorphism in the promoter/enhancer region of the FOXP3/Scurfin gene associated with type 1 diabetes. Immunogenetics, 2003. 55(3): p. 149-56.

161. Beier, U.H., et al., Histone deacetylases 6 and 9 and sirtuin-1 control Foxp3+ regulatory T cell function through shared and isoform-specific mechanisms. Sci Signal, 2012. 5(229): p. ra45.

162. Weir, G.C. and S. Bonner-Weir, Five stages of evolving beta-cell dysfunction during progression to diabetes. Diabetes, 2004. 53 Suppl 3: p. S16-21.

163. Porte, D., Jr. and S.E. Kahn, beta-cell dysfunction and failure in type 2 diabetes: potential mechanisms. Diabetes, 2001. 50 Suppl 1: p. S160-3.

164. Chakravarthy, M.V. and C.F. Semenkovich, The ABCs of beta-cell dysfunction in type 2 diabetes. Nat Med, 2007. 13(3): p. 241-2.

165. Bergman, R.N., D.T. Finegood, and S.E. Kahn, The evolution of beta-cell dysfunction and insulin resistance in type 2 diabetes. Eur J Clin Invest, 2002. 32 Suppl 3: p. 35-45.

166. Moynihan, K.A., et al., Increased dosage of mammalian Sir2 in pancreatic beta cells enhances glucose-stimulated insulin secretion in mice. Cell Metab, 2005. 2(2): p. 105-17.

167. Rousset, S., et al., The biology of mitochondrial uncoupling proteins. Diabetes, 2004. 53 Suppl 1: p. S130-5.

168. Banks, A.S., et al., SirT1 gain of function increases energy efficiency and prevents diabetes in mice. Cell Metab, 2008. 8(4): p. 333-41.

169. Robson-Doucette, C.A., et al., Beta-cell uncoupling protein 2 regulates reactive oxygen species production, which influences both insulin and glucagon secretion. Diabetes, 2011. 60(11): p. 2710-9.

170. Iino, S., et al., The antimetastatic role of thrombomodulin expression in islet cell-derived tumors and its diagnostic value. Clin Cancer Res, 2004. 10(18 Pt 1): p. 6179-88.

171. Dai, F.F., et al., The neuronal Ca2+ sensor protein visinin-like protein-1 is expressed in pancreatic islets and regulates insulin secretion. J Biol Chem, 2006. 281(31): p. 21942-53.

172. Huang, X., et al., The Identification of Novel Proteins That Interact With the GLP-1 Receptor and Restrain its Activity. Mol Endocrinol, 2013.

173. Wang, X., et al., Gene and protein kinase expression profiling of reactive oxygen species-associated lipotoxicity in the pancreatic beta-cell line MIN6. Diabetes, 2004. 53(1): p. 129-40.

67

174. Hardy, A.B., et al., Characterization of Erg K+ channels in alpha- and beta-cells of mouse and human islets. J Biol Chem, 2009. 284(44): p. 30441-52.

175. Basford, C.L., et al., The functional and molecular characterisation of human embryonic stem cell-derived insulin-positive cells compared with adult pancreatic beta cells. Diabetologia, 2012. 55(2): p. 358-71.

176. Kim, H., et al., Protein kinase Cbeta selective inhibitor LY333531 attenuates diabetic hyperalgesia through ameliorating cGMP level of dorsal root ganglion neurons. Diabetes, 2003. 52(8): p. 2102-9.

177. Wijesekara, N., et al., Beta cell-specific Znt8 deletion in mice causes marked defects in insulin processing, crystallisation and secretion. Diabetologia, 2010. 53(8): p. 1656-68.

178. Diao, J., et al., UCP2 is highly expressed in pancreatic alpha-cells and influences secretion and survival. Proc Natl Acad Sci U S A, 2008. 105(33): p. 12057-62.

179. Fu, A., et al., Loss of Lkb1 in adult beta cells increases beta cell mass and enhances glucose tolerance in mice. Cell Metab, 2009. 10(4): p. 285-95.

180. Wicksteed, B., et al., Conditional gene targeting in mouse pancreatic ss-Cells: analysis of ectopic Cre transgene expression in the brain. Diabetes, 2010. 59(12): p. 3090-8.

181. Chan, C.B., R.M. MacPhail, and K. Mitton, Evidence for defective glucose sensing by islets of fa/fa obese Zucker rats. Can J Physiol Pharmacol, 1993. 71(1): p. 34-9.

182. Price, N.L., et al., SIRT1 is required for AMPK activation and the beneficial effects of resveratrol on mitochondrial function. Cell Metab, 2012. 15(5): p. 675-90.

183. Sulaiman, M., et al., Resveratrol, an activator of SIRT1, upregulates sarcoplasmic calcium ATPase and improves cardiac function in diabetic cardiomyopathy. Am J Physiol Heart Circ Physiol, 2010. 298(3): p. H833-43.

184. Denton, R.M., Regulation of mitochondrial dehydrogenases by calcium ions. Biochim Biophys Acta, 2009. 1787(11): p. 1309-16.

185. Das, A.M., Regulation of mitochondrial ATP synthase activity in human myocardium. Clin Sci (Lond), 1998. 94(5): p. 499-504.

186. Rauchova, H., P.P. Kaul, and Z. Drahota, Activation of mitochondrial glycerol 3-phosphate dehydrogenase by cadmium ions. Gen Physiol Biophys, 1985. 4(1): p. 29-33.

187. Cardoso, C.M., et al., 4-Hydroxytamoxifen induces slight uncoupling of mitochondrial oxidative phosphorylation system in relation to the deleterious effects of tamoxifen. Toxicology, 2002. 179(3): p. 221-32.

188. Johansson, H., et al., Effect of fenretinide and low-dose tamoxifen on insulin sensitivity in premenopausal women at high risk for breast cancer. Cancer Res, 2008. 68(22): p. 9512-8.

189. Feil, R., et al., Ligand-activated site-specific recombination in mice. Proc Natl Acad Sci U S A, 1996. 93(20): p. 10887-90.

190. Feil, R., et al., Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains. Biochem Biophys Res Commun, 1997. 237(3): p. 752-7.

191. Liu, Y., et al., Tamoxifen-independent recombination in the RIP-CreER mouse. PLoS One, 2010. 5(10): p. e13533.

192. Sakurai, T., et al., Orexins and orexin receptors: a family of hypothalamic neuropeptides and G protein-coupled receptors that regulate feeding behavior. Cell, 1998. 92(4): p. 573-85.

68

193. Offield, M.F., et al., PDX-1 is required for pancreatic outgrowth and differentiation of the rostral duodenum. Development, 1996. 122(3): p. 983-95.

194. Swift, G.H., et al., An endocrine-exocrine switch in the activity of the pancreatic homeodomain protein PDX1 through formation of a trimeric complex with PBX1b and MRG1 (MEIS2). Mol Cell Biol, 1998. 18(9): p. 5109-20.

195. Chen, C. and E. Sibley, Expression profiling identifies novel gene targets and functions for Pdx1 in the duodenum of mature mice. Am J Physiol Gastrointest Liver Physiol, 2012. 302(4): p. G407-19.

196. Howitz, K.T., et al., Small molecule activators of sirtuins extend Saccharomyces cerevisiae lifespan. Nature, 2003. 425(6954): p. 191-6.

197. Minami, A., et al., Increased insulin sensitivity and hypoinsulinemia in APS knockout mice. Diabetes, 2003. 52(11): p. 2657-65.

198. Oriente, F., et al., Prep1 deficiency induces protection from diabetes and increased insulin sensitivity through a p160-mediated mechanism. Mol Cell Biol, 2008. 28(18): p. 5634-45.

199. Holst, J.J., The physiology of glucagon-like peptide 1. Physiol Rev, 2007. 87(4): p. 1409-39.

200. Wiederkehr, A. and C.B. Wollheim, Minireview: implication of mitochondria in insulin secretion and action. Endocrinology, 2006. 147(6): p. 2643-9.

201. Noda, M., et al., Switch to anaerobic glucose metabolism with NADH accumulation in the beta-cell model of mitochondrial diabetes. Characteristics of betaHC9 cells deficient in mitochondrial DNA transcription. J Biol Chem, 2002. 277(44): p. 41817-26.

202. Kennedy, E.D., P. Maechler, and C.B. Wollheim, Effects of depletion of mitochondrial DNA in metabolism secretion coupling in INS-1 cells. Diabetes, 1998. 47(3): p. 374-80.

203. Vetterli, L., et al., Resveratrol potentiates glucose-stimulated insulin secretion in INS-1E beta-cells and human islets through a SIRT1-dependent mechanism. J Biol Chem, 2011. 286(8): p. 6049-60.

204. Wu, Z., et al., Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell, 1999. 98(1): p. 115-24.

205. Uldry, M., et al., Complementary action of the PGC-1 coactivators in mitochondrial biogenesis and brown fat differentiation. Cell Metab, 2006. 3(5): p. 333-41.

206. Lehman, J.J., et al., Peroxisome proliferator-activated receptor gamma coactivator-1 promotes cardiac mitochondrial biogenesis. J Clin Invest, 2000. 106(7): p. 847-56.

207. Yoon, J.C., et al., Suppression of beta cell energy metabolism and insulin release by PGC-1alpha. Dev Cell, 2003. 5(1): p. 73-83.

208. Wild, S., et al., Global prevalence of diabetes: estimates for the year 2000 and projections for 2030. Diabetes Care, 2004. 27(5): p. 1047-53.

209. Yoshino, J., et al., Nicotinamide mononucleotide, a key NAD(+) intermediate, treats the pathophysiology of diet- and age-induced diabetes in mice. Cell Metab, 2011. 14(4): p. 528-36.

210. Maassen, J.A., et al., Mitochondrial diabetes: molecular mechanisms and clinical presentation. Diabetes, 2004. 53 Suppl 1: p. S103-9.

211. Baur, J.A. and D.A. Sinclair, Therapeutic potential of resveratrol: the in vivo evidence. Nat Rev Drug Discov, 2006. 5(6): p. 493-506.

69

212. Baur, J.A., et al., Resveratrol improves health and survival of mice on a high-calorie diet. Nature, 2006. 444(7117): p. 337-42.

213. Hubbard, B.P., et al., Evidence for a common mechanism of SIRT1 regulation by allosteric activators. Science, 2013. 339(6124): p. 1216-9.