University of Southern California Dissertations and Theses

Short term high fat diet (HFD) stimulates β cell proliferation through mTOR while the prolonged treatment induces β cell senescence via p27

(USC Thesis Other)

Short term high fat diet (HFD) stimulates β cell proliferation through mTOR while the prolonged treatment induces β cell senescence via p27

PDF

Download a page range

Download transcript

Copy asset link

Request this asset

Transcript (if available)

Content i
Short term High Fat Diet (HFD) stimulates β cell proliferation through
mTOR while the prolonged treatment induces β cell senescence via p27

By

Richa Aggarwal

A Dissertation presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR PHARMACOLOGY AND TOXICOLOGY)

May 2017

Copyright 2017 Richa Aggarwal
ii
Dedication

To my family

iii
Acknowledgements

There is a famous saying by F. Scott Fitzgerald, that “You don't write
because you want to say something; you write because you've got something to
say”.
Indeed, I’m here to thank a lot of people who made my journey through PhD
possible.
First and foremost, I would like to thank my advisor Dr. Stiles for accepting
me as a PhD student in her lab and giving me an opportunity to work on the β
cell project. I am grateful for her guidance and persistent support through my
project. She has been a great mentor whose invaluable suggestions opened my
mind to new possibilities and helped me grow as a research scientist all these
years. Her passion and commitment for science made my PhD experience very
productive and stimulating. Additionally, I cannot thank her enough for supporting
me by providing research assistantship for 4 years.
I want to express my sincere gratitude towards my Thesis Committee
Members Dr. Okamoto and Dr. Watanabe for their time, indispensable advice
and constructive criticism throughout the project. They have been constant
source of support and encouragement. I also want to thank Dr. Cadenas and Dr.
Li for their brilliant comments and suggestions during and after my oral qualifying
exam. Their insightful questions propelled my research in the right direction.
I am grateful to God for giving me the best bunch of friends here at USC,
especially Anketse Debebe. During the first couple of years of the program, my
friends in PIBBS program as well as at School of Pharmacy made tough times
iv
look easy. We shared classes and were together through good and bad grades. I
was super lucky to have lab mates who were extremely supportive and created a
very comfortable working environment. Ni Zeng, a senior graduate student, was
my mentor who taught me basic techniques in lab and how to handle mice. She
was our encyclopedia in lab and had answers to all our questions. I am really
grateful for her continuous support and assistance. I also want to thank my other
lab mates Anketse Debebe, Zhechu Peng, Lina He, Yang Li, Jingyu Chen,
Joshua Chen, Kelly Yang, Chengyou Jia, Indra Mahajan, Mengchen, Yating Guo
and Fan Fei for their unwavering support, stimulating discussions and
constructive feedback during lab meetings. Their love made our lab, my second
home.
My deepest gratitude goes to my family members- mom, dad, sister,
brother and in-laws for their constant love and support. Their unconditional love
made me a stronger person whenever I faced a setback. Thank you all for
believing in me and always wanting the best for me. Last but not the least, I want
to thank my husband, Rohit, who has been there for me in tough times. He has
always motivated me to strive better and kept me moving towards my goal. His
infinite belief in me and faithful support helped me go through these final stages
of PhD. Finally, I want to thank my daughter, Myra, for being my strength and
inspiration.

v
Table of Contents
Dedication .................................................................................................................................. ii
Acknowledgements ............................................................................................................... iii
List of Figures .......................................................................................................................... vii
List of Tables ............................................................................................................................ ix
Abstract ....................................................................................................................................... x
Chapter I: Overview of Obesity, Diabetes and β cell mass ......................................... 1
I-1 Obesity and Diabetes ................................................................................................................. 1
I-2 β cell development and regeneration ................................................................................. 2
I-3 Signaling pathways regulating β cell proliferation ......................................................... 3
I-4 Cell cycle regulators in pancreatic β cells .......................................................................... 6
I-5 Effect of high glucose and fatty acids on β cells ................................................................ 9
I-6 Significance ................................................................................................................................. 13
I-7 Main Hypothesis ....................................................................................................................... 13
Chapter II: High Fat Diet (HFD) induced effect on β cells ........................................ 15
II-1 Introduction and Rationale ................................................................................................. 15
II-1-1 β cells and their plasticity ............................................................................................................ 15
II-1-2 HFD as an obesity/ Diabetes model ........................................................................................ 16
II-2 Results- Role of CyclinD2 as a molecule that helps β cells to proliferate and
adapt as per the requirements ................................................................................................... 17
II-2-1 Glucose as a main driver for β cell proliferation (Figure 3)- ........................................ 18
II-2-2 Lipid and β cell proliferation (Figure 4)- .............................................................................. 20
II-3 Summary .................................................................................................................................... 22
Chapter III: Short term High Fat Diet (HFD) increases proliferation in β cells by
upregulating Cyclin D2 expression through mTOR signaling ............................... 24
III-1 Introduction and Rationale- .............................................................................................. 24
III-2 Results- ..................................................................................................................................... 28
III-2-1 β cell proliferation increases as early as 14 days after HFD treatment ................. 28
III-2-2 Cyclin D2 levels significantly increased in β cells after 2 month HFD treatment
leading to increased proliferation ........................................................................................................ 33
III-3 Discussion- ............................................................................................................................... 43
Chapter IV: Long term High Fat Diet (HFD) decreases proliferation in β cells by
down-regulating CyclinD2 expression and up-regulating p27 ............................. 48
IV-1 Introduction & Rationale- ................................................................................................... 48
IV-2 Results- ...................................................................................................................................... 52
IV-2-1 β cell proliferation decreases after 12-14 month HFD treatment ............................ 52
IV-2-2 Cyclin D2 levels significantly decrease while p27 levels increase after long term
HFD treatment ............................................................................................................................................... 56
IV-2-3 Chronic high levels of Triglycerides (TG) may be responsible for increased p27
and decrease in β cell proliferation. ..................................................................................................... 58
IV-2-4 Treatment with saturated fatty acids in vitro causes decreased viability,
proliferation and increased apoptosis ................................................................................................ 61
IV-3 Discussion ................................................................................................................................ 64
Chapter V: Overall Discussion .......................................................................................... 68
vi
Chapter VI: Future Directions .......................................................................................... 75
VI-1 To determine if mTOR is a fatty acid sensor ................................................................ 75
VI-1-1 Background & Rationale ............................................................................................................. 75
VI-1-2 Proposed Experiments ................................................................................................................ 77
VI-2 To determine if AKT activation can rescue the anti-proliferative effect of long
term activation of mTOR signaling on β cells ........................................................................ 78
VI-2-1 Background & Rationale ............................................................................................................. 79
VI-2-2 Proposed Experiments ................................................................................................................ 82
Chapter VII: Materials and Methods ............................................................................... 84
Bibliography ........................................................................................................................... 91

vii
List of Figures

Figure 1. Schematic diagram representing the effect of acute and chronic
hyperglycemia and hyperlipidemia on β cell proliferation and function leading to
Type 2 Diabetes
Figure 2. Schematic diagram showing various signaling pathways that are
activated by growth factors such as Insulin, IGF-1, HGF & PDGF and regulate β
cell proliferation
Figure 3. Correlation of cell proliferation with changes of plasma glucose levels
Figure 4. Correlation of cell proliferation with changes of plasma lipid levels
Figure 5. 14 day HFD feeding increased body weight and caused hyperlipidemia
in mice
Figure 6. Glucose intolerance and insulin resistance is seen in mice as early as
14 days on HFD
Figure 7. HFD significantly increased proliferation in β cells of mice fed for 14
days
Figure 8. Cyclin D1 and D2 levels do not change in mice after 14 day HFD
treatment in mice islets
Figure 9. 2 month HFD treatment cause increased body weight, hyperglycemia
and hyperinsulinemia
Figure 10. Glucose intolerance and insulin resistance is observed in mice after 2
month HFD treatment
Figure 11. High levels of plasma NEFA and triglyceride (TG) levels indicate
hyperlipidemia in mice treated with HFD for 2 months
Figure 12. HFD significantly increased proliferation in β cells of mice fed for 2
months
Figure 13. HFD feeding for 2 months significantly increased Cyclin D2 levels
Figure 14. In vitro treatment of INS-1 cell line with Palmitic Acid (PA) induces p-
S6K
Figure 15. Ingenuity Pathway Analysis (IPA) results showing significant
enrichment of the mTOR pathway in HFD islets
Figure 16. Rapamycin treatment abolishes HFD-induced β cell proliferation
Figure 17. In vitro treatment of INS-1 cell line with Palmitic Acid (PA) induces p-
ERK but does not mediate Cyclin levels via ERK pathway
Figure 18. HFD for 9-14 months decreases β cell proliferative potential and
subsequent islet area decrease
Figure 19. Blood Biochemical analysis of mice treated with HFD for 9 months
and 12-14 months
Figure 20. Glucose intolerance and insulin resistance after 9M and 14M of HFD
feeding
Figure 21. HFD feeding for 9M and 14M significantly decreased Cyclin D2 levels
Figure 22. Triglyceride (TG) levels significantly increase in HFD fed mice as
compared to NC control mice
Figure 23. Increased Triglyceride levels cause accumulation of p27 in β cells
after 14M HFD treatment causing decrease in proliferative potential
viii
Figure 24. Palmitic Acid (PA) decreases cell viability, growth and increases cell
apoptosis in vitro
Figure 25. Schematic diagram showing mTOR’s negative feedback signaling to
IRS and AKT

ix
List of Tables

Table 1. Diet composition table

x
Abstract
Hyperlipidemia and hyperglycemia are both hallmarks of Diabetes. β cells
are exposed to high levels of free fatty acids (FFAs) along with glucose during
Diabetes development. In this study, we focused on the exposure-dependent
effect of FFAs on β cells by using a High Fat Diet (HFD) model in mice and
Palmitic Acid (PA) treatment in INS-1 cell line. Both glucose and FFAs have been
earlier observed to have pro-proliferative effect upon short-term exposure and
can lead to cell death upon long-term exposure in β cells. However, the
molecular players modulating such effect are still unknown. Since β cell
proliferation is regulated by cell cycle proteins such as Cyclins, CDKs (Cyclin
dependent kinases) and CDKIs (Cyclin dependent kinase inhibitors), we
hypothesized that short term exposure of FFAs along with glucose will increase
cell proliferation while the long term will decrease cell growth via the cell cycle
regulatory proteins. To test this hypothesis, we fed mice with HFD for 14 days
and 2 months (short-term) and 9 & 14 months (long-term). Indeed, β cell
proliferation was significantly upregulated after short-term exposure, which was
accompanied by both hyperglycemia and hyperlipidemia. This β cell proliferation
was due to up-regulation of Cyclin D2 in mice islets. Our in vitro PA treatment
results further demonstrated that Cyclin D2 induction was regulated by the lipid
mediated activation of mTOR pathway, independent of AKT. RNA seq analysis of
the islets from HFD vs NC fed mice supported significant enrichment of the
genes involved in the mTOR pathway. Additionally, this result was confirmed in
xi
mice by using rapamycin (mTOR inhibitor) which abolished HFD mediated
proliferative effect.
Furthermore, long term HFD exposure decreased β cell proliferation.
Diabetic individuals experience decrease in β cell mass due to which they can no
longer maintain glucose homeostasis. In our long term HFD treatment model,
plasma triglycerides (TG) were significantly upregulated, along with decrease in
Cyclin D2 and increase in p27 levels in islets. In vitro treatment of INS-1 cells
with TG confirmed its effect on both Cyclin D2 and p27, suggesting a direct
impact of lipids on β cell proliferation and possible effect of TG on β cell
senescence.
Together, we found that HFD/lipids regulate β cell proliferation in an
exposure dependent manner via mTOR, Cyclin D2 and p27. Studies identifying
molecular regulators of β cell proliferation can contribute in targeted therapies
that can increase β cell mass in Diabetic individuals and alleviate Diabetes
symptoms.

xii
EMPTY PAGE
1

Chapter I: Overview of Obesity, Diabetes and β cell mass

I-1 Obesity and Diabetes

Obesity is associated with increased peripheral insulin resistance, which is
a major hallmark of Type 2 Diabetes pathophysiology. Obesity as an epidemic is
on the rise. As per the National Center for Health statistics, prevalence of obesity
was 36.5% among US adults during 2011-2014, with higher prevalence in older
adults. Along with genetic factors, sedentary lifestyle and caloric abundance are
the main causes for the increased incidence. Obesity is a major risk factor for
Diabetes. However, the relationship between obesity and diabetes is complex.
Insulin resistance refers to the increased insulin need in the peripheral
tissues such as adipose tissue, muscle and liver to enhance glucose uptake and
inhibit glucose output. In majority individuals, this increased insulin requirement is
met by increased insulin secretion by pancreatic β cells. Insulin (51 amino-acid
peptide) is a key hormone that regulates glucose homeostasis. Pancreatic β cells
secrete insulin in response to increased blood glucose. When released by β
cells, insulin signals glucose uptake by peripheral tissues such as muscle and fat
while suppresses gluconeogenesis in liver. It also stimulates conversion of
glucose to glycogen for storage in organs such as liver and skeletal muscles.
However, a subset of the population is unable to meet the increased insulin
demands and experience β cell dysfunction (decreased insulin secretion and cell
death) instead. This latter population manifests increasing glucose levels
(hyperglycemia) and is ultimately diagnosed with type II diabetes [1]. In other
2
words, in diabetes patients, the amount of insulin produced is not adequate for
the glucose levels, leading to hyperglycemia, the hallmark of diabetes. Therefore,
diabetes is a disease of β cells that cause imbalance between insulin needs and
its supply and hence maintaining β cell mass and function is crucial in
management of this disease.

Figure 1. Schematic diagram representing the effect of acute and chronic
hyperglycemia and hyperlipidemia on β cell proliferation and function
leading to Type 2 Diabetes.

I-2 β cell development and regeneration

Diabetic individuals experience loss of β cell mass and hence are unable
to maintain glucose homeostasis, leading to hyperglycemia. Understanding β cell
mass regulation is the key to understand diabetes, which is marked by either
near-absolute (type I) or relative (type II) β cell deficiency. Hence, islet
3
regeneration has been the main focus of the research studies aiming at
replenishment of β cells in diabetic patients.
During embryogenesis, β cells arise from the stem/progenitor cells in the
gut endoderm. Genetic tracing experiments indicate that the endocrine and
exocrine cells of the pancreas are derived from endodermal cells expressing
transcription factors Pdx1[2] and Ptf1a [3]. Late fetal gestation period
experiences fastest expansion of β cell population, with nearly doubling of the
cells each day starting from 16th day post conception in rats [4]. Growth of β cell
mass still occurs in the neonatal rodents mainly due to neogenesis from the duct-
like cells as indicated by BrDU labeling data [5], with a minor contribution from
the mitotic activity of the existing β cells [6]. β cells proliferate at a very low rate
during adulthood, and this rate gradually declines as the organism ages. Studies
in rats have indicated that in 30-100 day old mice, approximately 1-4% β cells
proliferate [7], while, long-term BrDU labeling experiments have shown that only
1 in 1400 (approx.) β cells replicate per day in a one-year-old mice, corroborating
to the very slow β cell turnover in adults [8].

I-3 Signaling pathways regulating β cell proliferation

Although very low proliferation rate is seen in adults under normal
physiological conditions, β cell mass is seen to expand during pregnancy and
obesity to compensate for the increased insulin demand and during pancreatic
injury and repair [9]. While the exact mechanism for such effect is still unclear,
linear tracing studies have indicated that pre-existing β cells are the source of
4
replication in adult islets [10]. The signals that motivate β cells to replicate are not
clearly understood, though glucose and various growth factors have been
observed to exhibit a mitogenic effect.
Growth factors (such as PDGF (platelet derived growth factor), HGF
(Hepatocyte growth factor), insulin and IGF-1 (Insulin-like growth factor 1)),
incretins, glucose and amino acids have been shown to influence β cell
proliferation. Although there are diverse mechanisms that induce proliferation in β
cells, insulin-dependent signaling pathways play an important role in regulating
insulin secretion and β cell growth, indicating that β cells are not only the source
of insulin production but also its major target. Mitogens such as Insulin and
insulin like growth factor-1 (IGF-1) bind to their receptors (receptor tyrosine
kinases) on the cell surface and activate downstream proteins such as IRS
(insulin receptor substrate) and GAB-1 to mediate β cell survival and
proliferation. Deletion studies where insulin receptor (IR) is lost specifically in β
cell specific showed that glucose-stimulated insulin secretion is inhibited [11]. In
these mice, reduced β cell mass is also observed with aging. Mice lacking IGF-1
receptor are defective in insulin secretion in response to glucose [12, 13].
Hepatic growth factor (HGF) has also been reported to influence islet mass. HGF
receptor deletion from β cells causes smaller islets and enhanced apoptosis [14]
while over-expression leads to increased proliferation [15].
Phosphoinositide 3-kinase (PI3K)/AKT signaling pathway plays a major
role downstream of insulin/IGF-1 and HGF signaling cascades to promote β cell
proliferation and function [16, 17]. Mice expressing constitutively active form of
5
AKT showed larger islets and reduced rate of apoptosis after streptozotocin
(STZ) treatment [18]. Additionally, study by Fatrai et al has indicated that AKT
over expression causes increased β cell proliferation by regulating Cyclin D1, D2
and p21 levels, giving insights on the molecular mechanisms responsible for the
effect [19]. AKT regulates mTOR (Target of Rapamycin) activity by
phosphorylating TSC2 (tuberous sclerosis complex 2) and positively influencing
the pathway [20]. mTOR responds to growth factors and nutrient signals and is
essential for β cell growth and proliferation. Studies indicate that mTOR is a part
of two different complexes-mTORC1 and mTORC2. mTORC1 complex contains
Raptor and the G protein β-subunit-like protein (GβL) and activates protein
translation regulators such as ribosomal S6 kinase (S6K), eukaryote initiation
factor 4E-binding protein 1 (4EBP1), and eukaryote initiation factor 4E. This
complex is sensitive to rapamycin, an antifungal compound produced by
bacterium Streptomyces hygroscopicus. mTORC2 on the other hand, contains
Rictor, is rapamycin insensitive and it phosphorylates/activates AKT at Ser473
[21-23]. Several studies have provided evidence for mTOR signaling in β cell
mass/proliferation using genetically modified mice models. S6K
-/-
mice are
significantly smaller at birth as compared to the wild type [24] with smaller
pancreatic islets [25] leading to hypoinsulinemia and glucose intolerance. TSC2
is a negative regulator of mTORC1 signaling. Deletion of TSC2 specifically in β
cells results in activated mTOR signal and increased β cell proliferation, β cell
size and hyperinsulinemia [26, 27]. Treatment with rapamycin (which inhibits
mTOR) in pregnant mice greatly reduced β cell proliferation, pancreatic islet
6
content and islet size [28]. Furthermore, study by Balcazar et al indicated that
mTOR regulates β cell proliferation by modulating Cyclin D2, D3 and cdk4 levels.
They also showed that mTOR controlled Cyclin D2 synthesis and stability [29].
All these studies clearly indicate the important role of mTOR signaling pathway in
maintaining β cell proliferation and size.

Figure 2- Schematic diagram showing various signaling pathways activated
by growth factors such as Insulin, IGF-1, HGF & PDGF and regulate β cell
proliferation.

I-4 Cell cycle regulators in pancreatic β cells

Replication of pre-existing β cells is the primary mechanism in adults that
maintains β cell mass and compensate for the increased insulin requirement
under physiological condition [6, 10]. Cell cycle progression in β cells is
controlled by 3 major groups of proteins- cyclins, cyclin-dependent kinases
7
(CDKs) and cyclin dependent kinase inhibitors (CDIs). Cyclin Ds are proteins that
regulate G1/S phase cell cycle transition by interacting with cdk4 or 6, in
response to mitogenic activation. β cells contain three D-Cyclins – D1, D2 and
D3, out of which Cyclin D1 and D2 are mainly important for cell proliferation.
According to Georgia and colleagues, islet size is markedly reduced in Cyclin D2
null mice, along with reduced serum insulin levels [30]. Kushner JA. et al further
demonstrated the reduction of β-cell replication is exacerbated in Cyclin D1 +/-
D2 -/- mice. These mice had abnormally small islets, experienced severe
hyperglycemia and death by 4 months of age. On the other hand, Cyclin D2-/-
mice had normal islet mass at 3 months but had 70% reduction at 9-12 months
indicating the role of Cyclin D2 in postnatal islet growth [31]. In addition, the
Stewart group showed that over-expression of Cyclin D1/cdk4 using adenovirus,
leads to dramatically increased proliferation rate of β-cells [32]. Together these
studies support the essential role of Cyclin D1 and D2 in contributing to the
replicative potential of β cells.
Inhibitors of the cell cycle that prevent formation of active cyclin/CDK
complexes are divided into two families- the INK4/ARF family composed of
p16
INK4a
, p15
INK4b
, p18
INK4c
and p19
INK4d
; and the CIP/KIP family composed of
p21
Cip1
, p27
Kip1
and p57
Kip2
. p16ink4a is a well-known marker of aging and has
been seen to cause age-dependent loss in β cell regenerative capability. As per
the study by Krishnamurthy et al, p16
INK4a
expression was 14-fold higher in older
mice (64-80 weeks old) when compared to levels in 12-14 weeks old mice. While
overexpression of p16
INK4a
in young islets significantly reduced β cell
8
proliferation, its deletion in old β cells rescued the age-dependent proliferation
defect [33]. These data strongly indicate the crucial role of p16
INK4a
in β cell
proliferation.
Furthermore, a lot of recent studies have related the non-progression of
cell cycle in β cells to the expression levels of p27. P27 is a marker of
senescence which has been shown to accumulate in terminally differentiated β
cells during embryogenesis [34]. Study by Georgia and Bhushan, 2006 showed
that by deleting p27 in the mice during embryogenesis can increase islet mass,
cause hyperinsulinemia and prevent hyperglycemia after STZ (β cell toxin)
treatment in these mice as compared to the control. This study indicated that
deleting p27 allows the cell to enter the cell cycle and promotes their replication
[34]. Additionally, in two well established Type 2 Diabetes genetic models (Insulin
receptor substrate2 deletion (Irs2-/-) and leptin receptor deletion (Lepr-/-)) p27
was shown to be accumulated in β cells. While overexpression of p27 (Cdkn1b
gene) caused severe diabetes in these mice, its deletion prevented
hyperglycemia through increasing islet mass and serum insulin [35]. These
results highlight the role of p27 in maintaining β cell mass and its involvement in
T2D pathogenesis. In depth analysis of role of p27 at different stages of
development by Latif Rachdi et al. indicated that, overexpression of p27 in adult
life (8 weeks-26 weeks) did not have significant effect on β cell mass and
proliferation [36]. On the other hand, its overexpression after birth or during the
first 4 weeks of postnatal life resulted in glucose intolerance and reduced β cell
9
mass. These observations suggest that p27 plays a major role in regulating β cell
mass during early developmental stages.
Overall the above evidences indicate that the cell cycle in β-cells can be
modulated through the basic regulatory machinery similar to non-islet tissues.
The levels of Cyclins/CDKs and CDKIs and their interaction in the cell during
replication decides whether or not the cell goes into division and hence makes
them an attractive target in regulation of β cell replication.

I-5 Effect of high glucose and fatty acids on β cells

Diabetic individuals are exposed to chronic high levels of glucose and fatty
acids. A phenomenon called “glucotoxicity” describes the negative effect chronic
elevated levels of glucose on β cell function. Glucose stimulates insulin
production and release under normal conditions but prolonged hyperglycemia
causes excessive β-cell degranulation and eventual exhaustion. Various studies
in the past have documented the effect of glucose on β cells. Short term glucose
infusion (24-48hours) studies in 3-month old rats have indicated increase in β cell
number due to β cell neogenesis, hypertrophy and hyperplasia in glucose infused
rats as compared to controls [37, 38]. Similar hyperplasic effects of glucose have
been observed in human islets, which when exposed to 33.3mM glucose showed
increased number of replicating β cells (Ki67 positive cells) as compared to
5.5mM glucose treated islets [39]. These studies support the mitogenic effect of
glucose when exposed for short duration.
10
Glucose can mediate this mitogenic effect directly or via insulin. The
glucose induced mitogenic effect in β cells is mediated by Glucokinase (GK), an
enzyme involved in glycolysis. GK converts glucose to glucose-6-phosphate and
hence acts as a glucose sensor that can regulate insulin secretion in β cells [40].
A study reported that haploinsufficiency of GK (GK
+/-
) in β cells causes impaired
insulin secretion [41]. The same group later demonstrated that when fed High fat
diet (HFD) for 20 weeks, GK
+/-
mice have decreased β cell replication and
insufficient increase in β cell mass as compared to wild type counterparts [42].
These results support the idea that GK is essential for compensatory increase in
β cell mass in response to insulin resistance associated hyperglycemia.
Additionally, glucose-stimulated insulin released by β cells modulates intracellular
signaling pathways that promote proliferation in β cells. IRS-2 (Insulin receptor
substrate 2) is one of the major substrates of insulin signaling. A study by Kubota
et al has shown that the IRS-2 KO mice develop insulin resistance, have
significant reduction in β cell mass followed by development of Diabetes [43].
Furthermore, increased expression of IRS2 in the β cells prevented Diabetes in
IRS
-/-
mice, obese mice and Streptozotocin (β cell toxin) treated mice, at least
partially by promoting β cell growth, survival and insulin secretion [44]. Together,
these results support that glucose mediates its direct effect through GK to
facilitate β cell proliferation but indirectly through IRS-2 to promote β cell growth
and compensation under insulin resistance conditions.
While short-term hyperglycemia appears to induce proliferation of β cells
and secretion of insulin, excessive glucose decreases insulin expression by
11
decreasing its gene expression [45]. It also causes mitochondrial dysfunction &
ER stress; all factors contributing to β-cell dysfunction and loss seen in diabetic
patients [46].
“Lipotoxicity”, on the other hand, is often used to refer to the effect of free
fatty acids (FFAs) on cells in an insulin resistance setting. FFAs can have both
beneficial and detrimental effects [47] depending on the duration, coexistence
with glucose and the kind of FFA. While 4-day treatment of cultured human islets
with saturated fatty acids (such as palmitate) had pro-apoptotic effect,
unsaturated fatty acids (such as oleate) were seen to be protective against
palmitate and glucose-induced β-cell death [48]. Studies related to conditions
marked with acute hyperlipidemia such as pregnancy, have indicated that β cells
increase their mass and insulin secretion in response to increased insulin
demand in presence of high FFA conditions [49-51]. Additionally, in vitro studies
with rat islets have demonstrated higher BrDU incorporation by several folds in
palmitic acid treated islets versus controls indicating higher cell replication in
presence of fatty acids [52, 53]. While short-term exposure of FFAs is
proliferative, long-term exposure is deleterious for the β cells. Prolonged
exposure to FFAs causes decreased insulin transcription, impaired glucose-
induced insulin secretion (GSIS) and finally, β cell apoptosis [54]. Studies have
also found decreased β cell mass and higher rate of apoptosis as indicated by
BrDU incorporation and immunohistochemistry analysis in diabetic rats versus
the lean controls [45, 55]. A recent study by Pascoe et al indicated that free fatty
acids (such as palmitic acid) inhibit glucose-stimulated proliferation by
12
upregulating cell cycle inhibitors p16 and p18, both in vivo and in vitro, while
having no effect on Cyclin D2 expression [56].
Even though, prolonged glucose and fatty acids exposure separately have
negative impact on β-cell proliferation and insulin secretion, researchers believe
that it’s the combination of both that makes it worse [46]. This mechanism is
referred to as “Glucolipotoxicity”. Under normal glucose concentrations, fatty
acids are transported into the mitochondria by an enzyme called carnitine-
palmitoyl transferase-1 (CPT-1) and oxidized. But elevated glucose causes
increase in the formation of malonyl CoA via citrate, which inhibits CPT-1. This
results in accumulation of long chain acyl CoA (LC-CoA) esters in the cytosol
[57], which further increases ceramide and triglyceride levels in the cell.
Accumulation of ceramide causes inhibition of anti-apoptotic protein Bcl-2 and
down-regulation of IRS-1/2 signaling [58] leading to decreased β-cell proliferation
and cell death.
All this data together indicates that both glucose and fatty acids can act as
either growth-promoting or growth-deteriorating factors under different
physiological conditions affecting β cell mass and hence contributing to pathology
of diabetes. Although the aforementioned studies have considered the effect of
fatty acids on proliferation, the studies have been done in glucose rich conditions,
thereby making it harder to point out glucose independent effect of fatty acids.
Thus, we in this study are aiming to elucidate the effect of fatty acids
(independent of glucose) on β cell proliferation and mass, and also determine the
downstream cell cycle regulators that might be responsible for the effect.
13

I-6 Significance

29.1 million (9.3%) adults and children of the US population currently have
been diagnosed with diabetes, according to the American Diabetes Association
2014 fact sheet. Diabetes is a complex metabolic disease that not only causes
metabolic dysfunction but also affects the quality of life of an individual. It is also
the seventh leading cause of death in US individuals.

I-7 Main Hypothesis

β cells are primarily insulin producing cells in pancreatic islets, responsible
for maintaining glucose homeostasis. Although, in adults they have a very slow
turn-over rate, β cell mass has been seen to expand during pregnancy, obesity
and insulin resistance to compensate for the increasing insulin needs. Failure to
do so leads to pathology of type II diabetes. Type II diabetes patients are
exposed to high levels of glucose and free fatty acids (FFAs)/lipids. Studies have
indicated that glucose and FFAs/lipids can have either pro or anti-proliferative
effect on β cells. Acute exposure is shown to have proliferative effect leading to
increased islet mass and hence increased compensatory insulin secretion. On
the other hand, chronic high exposure to glucose and fatty acids/lipids causes
decrease in β cell proliferation and enhances the rate of cell apoptosis. However,
the exact molecular mechanism by which HFD causes its pro or anti-proliferative
effect is still debatable. Therefore, in this study we want to investigate the
early (acute) and delayed (chronic) effects of glucose and fatty acid/lipid
14
exposure on pancreatic β cells and identify key molecular components that
regulate β cell proliferation and death. In order to achieve our goal, we would
expose mice to high glucose and lipid environment by feeding them High Fat Diet
(HFD) for short (14 days and 2 months) and long duration (9 months and 14
months). Furthermore, to evaluate the effects of lipids (in presence of
physiological levels of glucose), the data will be supported by in vitro treatment of
INS-1 cell line (rat insulinoma) with fatty acids and triglycerides (TG).
Given the unique role of β-cells in diabetes studies, understanding β-cell
regulation will provide important directions for β-cell regeneration and potential
treatment approaches in type II diabetes patients.

15
Chapter II: High Fat Diet (HFD) induced effect on β cells

II-1 Introduction and Rationale

II-1-1 β cells and their plasticity

Pancreatic β cells, present in the islets of Langerhans, are responsible for
dynamically maintaining glucose homeostasis in the body. Adult β cells are
known to have a very low turn-over rate [7, 8]. It has been observed that the
maximum increase in the β cell pool occurs during late fetal gestation and the
neonatal period [4, 5]. Although various models of β cell regeneration and
replenishment after damage/injury have been proposed such as presence of
adult pancreatic stem cells in epithelium of pancreatic ducts [59-61], trans-
differentiation of pancreatic acinar cells [62] and islet-neogenesis [7, 59, 63],
researchers are now increasingly accepting the theory of self-duplication of
existing adult β cells for increased β cell mass [10, 30]. Any injury or loss in β
cells during adulthood is recovered by division of the already existing β cells,
depending on the age of the individual. Younger mice have been reported to
respond better to regeneration models such as exposure to High fat diet (HFD)
and STZ (Streptozotocin- β cell toxin) treatment and have higher β cell
proliferation rates as compared to the older counterparts [64]. These results
indicate that even though there is slow turn-over of β cells in healthy individuals,
in response to injury or damage, β cells have the ability to self-duplicate and
increase their number to compensate. Diabetic models also support this
observation, wherein during the pre-diabetic phase when insulin resistance starts
16
developing, an increase of β cell mass and function is observed to compensate
for the increasing demand. These findings collectively support the adaptive
replicative plasticity in adult β cells.

II-1-2 HFD as an obesity/ Diabetes model

Hyperglycemia and hyperlipidemia are major hallmarks of obesity and
Diabetes. Both glucose and fatty acids/lipids act as nutrient sources and have
been observed to influence β cell proliferation and mass under obesity and
Diabetic conditions. Various studies have suggested the effect of glucose and
lipids on β cells where short term exposure is beneficial and pro proliferation,
while prolonged exposure is detrimental and causes β cell death, referred to as
Glucolipotoxicity. In vivo, hyperlipidemia develops together with hyperglycemia
conditions in individuals as well as in rodent experimental models. High Fat diet
(HFD) is a lipid-rich diet that exposes mice to higher fat concentrations and has
been accepted to be a relevant model of diet-induced obesity/Diabetes [65, 66].
HFD studies done so far have utilized different fat content (ranging from 30-60%)
and feeding duration, which has led to differences in interpretation of the roles
that lipid may play in β cell growth. While short term feeding studies such as the
1 week HFD study by Stamateris et al [67], using 60% fat in HFD, shows
hyperglycemia but no changes in free fatty acids (FFAs) associated with
induction in β cell proliferation, long term study (1 year HFD treatment) by Sone
et al (60% fat) [68] indicate induction of hyperglycemia and hyperlipidemia (FFAs
and triglycerides) along with reduction in β cell proliferation and mass.
17
Additionally, 12 month long HFD study by Hull et al [69] utilized three different fat
content in mice diet (15%, 30% and 45%) and reported fat-dependent increase in
β cell mass associated with loss in β cell function. In this study, they observed no
hyperglycemia and did not measure plasma FFA levels but suspected that high
fat concentration due to the diet might be impacting β cells. Collectively, all these
studies indicate early induction of hyperglycemia in mice with HFD feeding
followed by hyperlipidemia and changes in β cell mass & proliferation. However,
it has been difficult to separate the effects of lipids independent from that of
glucose. Moreover, there is no study that has looked at longitudinally the effects
of high fat on β cells. In an effort to address a lipid specific effect on β cells, we
decided to perform detailed analysis on mice fed HFD for different durations to
evaluate the dynamic changes of glucose, lipid and β cell growth. Here we fed
mice with HFD for 14 days, 2 months, 9 months and 14 months; (where we
considered 14 days and 2 months as short term treatment and 9 months and 14
months as long term treatment), in order to study the dynamic/longitudinal
changes in plasma lipid and glucose and relate it to β cell proliferation and cell
death.

II-2 Results- Role of CyclinD2 as a molecule that helps β cells to proliferate
and adapt as per the requirements.
Cyclins Ds, along with their partner cyclin dependent kinases, regulate the
G1 phase of the cell cycle and help in progression. Three Cyclin Ds have been
identified in rodent models- Cyclin D1, D2 and D3 and vary in their expression
18
levels. In mice, Cyclin D2 has been recently identified as a molecule that plays a
critical role in adult (postnatal) β cell proliferation [31, 67, 70]. Cyclin D2 pairs
with cdk4 (cyclin dependent kinase 4) and helps in promoting G1-S phase
transition during cell division.
In order to identify and segregate the roles of glucose and fatty acids/lipids
on β cell proliferation, we analyzed plasma glucose, fatty acids and Triglyceride
(TG) levels over the duration of HFD treatment along with the IHC staining
(Ki67/BrDU and Cyclin D2) of the pancreatic islets at the end of the treatment
period.

II-2-1 Glucose as a main driver for β cell proliferation (Figure 3)-

In our study, we analyzed plasma glucose levels every week along with β
cell proliferation and levels of cyclin D2 at the end of each study i.e 14 days, 2
months, 9 months and 12-14 months. Our short term HFD treatment conditions
(14 days and 2 months) showed increased β cell proliferation, as seen by
upregulated Ki67 expression. Consistent with previous reports [68], long term
HFD feeding for 9 months and longer does not induce the proliferative response
in β cells. Our data actually indicated a loss of proliferation potential in mice fed
HFD for 9 and 14 months (Fig 3). Mice fed HFD for 2 months developed
hyperglycemia. In this group of mice, we observed approximately 13-fold
increase in β cell proliferation as indicated by Ki67 staining. This high Ki67
staining index correlated with significantly higher levels of Cyclin D2 in β cells of
these HFD mice group. As shown in figure 3, hyperglycemia is also most
19
significantly induced in this group of mice (2 months HFD) among the 4 feeding
groups, which corresponds with significant increase in Cyclin D2 levels in the β
cells of these mice. This observation is in accordance with recently published
studies that indicate that glucose might act as a mitogen driving cyclin D2
expression and hence proliferation in β cells [71, 72].
Figure 3. Correlation of cell proliferation with changes of plasma glucose
levels. Summary of
the relationship
between proliferation
(ki67), Cyclin D2 and
plasma fasting
glucose levels after
14 days, 2 months, 9
months and 14
months of HFD
feeding; the bars are
depicted as fold
change (log2) in HFD as compared to NC.

High fat diet feeding for 9 months and longer led to significant
downregulation of Cyclin D2 in mouse islets concurrent with lack of observable
difference in proliferation between NC and HFD groups as indicated by Ki67
staining. In these mice, hyperglycemia is also reduced, supporting the observed
20
correlation between glucose and cell proliferation in the 2 months HFD feeding
group. Together, these data indicate a pro-proliferative effect of glucose via
cyclin D2 on β cells that is lost with prolonged feeding of HFD.

II-2-2 Lipid and β cell proliferation (Figure 4)-

While 2 month HFD feeding leads to hyperglycemia and the correlated
increase of β cell growth, glucose levels did not correlate with cell growth in the
shorter (14 day HFD group) and longer (9 months and 14 months HFD groups)
groups. In these groups, fatty acids and TG appears to have a role in β cell
growth. Our results indicate that NEFA (non-esterified fatty acids) was
significantly elevated only in the 14 day HFD treatment group (Fig 4). In these
mice, increased Ki67 index did not correlate with hyperglycemia. Ki67 index was
3-fold higher in these mice with an insignificant increase of Cyclin D2 expression.
However, fasting glucose was not high but rather moderately reduced in these
mice. In this feeding group, plasma free fatty acid (NEFA) levels are 1.5-fold
higher in the HFD fed mice of this group indicating a potential role of free fatty
acids in sustaining β cell proliferation that is overridden later (in the 2Month HFD
group) by glucose during the hyperglycemia stage. This effect of NEFA is
corroborated by previously published studies that have indicated the pro-
proliferation effects of fatty acids on β cells [49, 53]. This effect of fatty acids is
unlikely to be mediated by cyclin D2 as cyclin D2 levels is not altered in the 14
day HFD feeding group.
21
Figure 4. Correlation
of cell proliferation
with changes of
plasma lipid levels.
Summary of the
relationship between
proliferation (ki67),
Cyclin D2, plasma fed
NEFA (non-esterified
fatty acids) and plasma
fed TG (triglycerides)
levels after 14 days, 2 months and 14 months of HFD feeding; the bars are
depicted as fold change (log2) in HFD as compared to NC.

Previous studies have reported that older mice do not respond to the
mitogenic stimuli [33]. In the 9 months HFD fed group, high plasma glucose was
unable to induce the expression of cyclin D2 or increase β cell proliferation (Fig
3). These observations suggest that other signals may interfere with the glucose
induced β cell growth and regulate β cell, cell cycle. In the 9 month HFD feeding
group, decreased β cell proliferation and cyclin D2 levels correlated with higher
levels of triglyceride (TG). A 1.5 fold increase of fasting TG is observed in the 9
month-HFD feeding group concurrent with approximately 1.5 fold decrease in
cyclin D2 levels. A lack of response of β cell proliferation is also observed even
22
though the mice are still moderately hyperglycemic at this time (Fig 3). It was
reported that older mice lose mitogenic stimulation as they undergo senescence
[68]. Therefore, we hypothesize that long-term exposure of high levels of TG may
play a role in β cell senescence and contribute to the loss of proliferation
response in the 9 and 14 month HFD feeding groups.

II-3 Summary

HFD model is a diet-induced obesity model that has been widely accepted
to study the impact on β cells during the development of insulin resistance,
obesity and Diabetes in mice. Although several studies have been published with
varied fat content in the HFD and varied lengths of HFD feeding, no study has
longitudinally followed the effect of HFD on β cells using both short-exposure and
prolonged feeding time points. Therefore, in our study we fed mice with HFD
(60% fat) for 14 days, 2 months, 9 months and 14 months. Our data indicates
time-dependent effect of HFD on β cell proliferation. We identified glucose as the
main driver for β cell proliferation at an early time point i.e 2 months with
significant increase in Ki67 positive β cells and upregulated Cyclin D2 levels
while TG to be mainly responsible for loss of β cell proliferation and induction of
cell senescence after 9 and 14 months HFD treatment. We further investigated
the signaling pathways & molecular mechanisms that might be responsible for
the HFD mediated glucose-induced pro-proliferation effect and TG facilitated loss
of β cell growth and induction of senescence in this study.
23
In summary, in this study we corroborated previous studies that have
shown early adaptive induction of β cell proliferation and loss of β cell
proliferative potential after long-term HFD treatment in mice and further identified
major mechanisms that might be responsible for such observation.

24
Chapter III: Short term High Fat Diet (HFD) increases proliferation in β cells
by upregulating Cyclin D2 expression through mTOR signaling

III-1 Introduction and Rationale-

Pancreatic β cells, located in islets of Langerhans, are responsible for
sensing plasma glucose levels and regulating its uptake by adipose tissue and
muscle, by releasing Insulin. Diabetes results from loss of β cell mass and
function and current therapies are limited to insulin replacement therapy, whose
levels are not well regulated by changes of plasma glucose. Major efforts are
focused on improving β cell numbers in diabetic patients by transplantation with
limited success. The use of therapeutic agents that can activate cell proliferation
in either pre-existing β cells or induce differentiation/trans-differentiation to form
new β cells has become a sought-after goal for diabetes researchers. To achieve
this goal, studies are needed to understand specific molecular pathways that can
be targeted to stimulate β cell proliferation and contribute to β cell regeneration
therapy.
β cell mass increases during embryogenesis (late fetal gestation) and has
a very limited turn-over rate throughout the life span of the individual [8].
However, compensatory increase in the β cell mass in adults does take place
under conditions such as Pregnancy, obesity or insulin resistance. Generally,
obesity and insulin resistance precedes type II diabetes wherein β cells respond
by increasing their mass and insulin secretory capacity to maintain euglycemia.
Consistent with this notion, increase in β cell volume has been seen in obese
25
non-diabetic patients [73], suggesting that adult β-cells have the plasticity to
regenerate their own mass.
Recent studies have shown that nutrients such as glucose, amino acids
and fatty acids can induce β cell proliferation [74, 75]. Under conditions such as
obesity and Diabetes, glucose and free fatty acids (FFAs) are the most important
drivers that determine β cell equilibrium as hyperglycemia and hyperlipidemia are
major hallmarks of its pathology. Both glucose and fatty acids have acute and
chronic effects on β cell mass and function. While acute exposure to high
glucose and lipids increases β cell viability and function, the chronic exposure is
found to be detrimental and causes β cell damage (glucolipotoxicity) [46, 47].
In this study, we concentrate on the acute mitogenic effects of glucose
and FFAs/lipids on β cell proliferation. Research published in the past has
established the pro-proliferative effect of glucose and FFAs on β cells. Short term
exposure (24-48 hours) to glucose in β cells have been reported to enhance β
cell numbers in young rats infused with glucose [37, 38]. Similar hyperplastic
effect of glucose has also been shown in human islets, wherein islets exposed to
33.3mM glucose showed increased number of replicating β cells (Ki67 positive
cells) as compared to 5.5mM glucose treated islets [39]. Additionally, studies
looking at the effect of FFAs on β cells have shown that FFAs induce glucose-
stimulated insulin secretion (GSIS) by β cells during insulin resistance conditions
[76]. Study by Milburn et al, has implied that rat islets exposed to FFAs (oleate
and palmitate) increase insulin secretion by two folds and BrDU incorporation by
3.2 folds, indicating higher β cell proliferation [53]. Moreover, palmitate and
26
oleate levels increase several folds during pregnancy [50, 51]. According to a
study mimicking pregnancy conditions by using prolactin in the ex vivo culture
medium, by Brelje et al, palmitate induced up to 2.5-fold increase in insulin
secretion and 3-fold increase in cell proliferation in cultured rat islets. While
oleate had a little effect on insulin secretion, it had a significant impact on β cell
proliferation as seen by BrDU labeled insulin positive cells [49]. Together these
results suggest that in vitro and ex vivo treatment with glucose and fatty acids
such as palmitate cause increase in β cell proliferation. Our data supports these
results, wherein we observe significant increase in Ki67 positive β cells after 2
months of HFD feeding in mice. These HFD fed mice experience significantly
higher levels of glucose and plasma NEFA (non-esterified fatty acids) as
compared to their NC counterparts.
In this study, we aim to use a physiologically relevant Diabetes model of
High fat Diet (HFD) feeding, in which β cells will be exposed to high glucose and
fat/lipid environment. HFD allows the development of insulin resistance, obesity
and adaptive islet compensatory response in stages similar to what is seen in
Diabetic individuals [67-69, 77, 78]. Using this model, we want to study HFD-
induced β cell proliferative responses at 2 different HFD feeding time points (14
days and 2 months) together with the accompanying changes in plasma glucose
and fatty acid levels.
Additionally, we want to investigate the downstream signaling
mechanisms responsible for the proliferation effect. Although many studies have
established the role of glucose in driving several mitogenic signaling pathways in
27
β cells including PI3K/AKT, mTOR and extracellular signal-related kinase (ERK),
effect of fatty acids/lipids on these pathways is not clearly elucidated. Some
studies have been published demonstrating the effect of palmitic acid (PA) (most
abundant FFA in blood) on AKT but they have shown inconsistent results. While
some studies have reported that 24-hour exposure of PA causes AKT activation
in INS-1 cells in dose-dependent manner [79], others showed that PA reduces
active AKT and induces apoptosis after 24-48 hour treatment [80, 81]. These
discrepancies appear to result from the different glucose levels used for the
different studies. Furthermore, the effect of fatty acids/lipids on MAPK/ERK and
mTOR pathway in β cells has not been addressed. Hence, in this study we will
address the effect of PA using INS-1 cell line. We will keep a 6mM glucose
concentration (physiological levels) to match physiological glucose levels in the
plasma. We believe these treatment conditions would better represent the effect
of fatty acids on β cells and would provide a clearer picture of its effect on various
signaling pathways. We will use this condition to identify the pathways that are
affected by fatty acids (in presence of glucose) and contributes to increased β
cell proliferative potential.
β cell proliferation is regulated by proteins in the cell cycle machinery.
These include Cyclins, Cyclin dependent kinases (Cdk) and Cdk inhibitors
(CDKI). Particularly, proteins that control the G1/S cell cycle transition have been
reported to play an important role in replication of β-cells. These include D cyclins
such as Cyclin D1 and D2, which associate with Cdk4/6 to make a complex that
phosphorylates and inactivates RB (Retinoblastoma tumor suppressor protein).
28
This phosphorylation causes the dissociation of elongation factor, E2F from RB,
which then activates the proteins required for the S phase. Several studies have
emphasized the role of Cyclin D1, D2, p27 and p53 in regulating β cell cycle [30-
32, 34, 82].
Our aim in this study is to elucidate short-term effect of high fat diet on β
cell proliferation and evaluate the downstream signaling pathways.

III-2 Results-

III-2-1 β cell proliferation increases as early as 14 days after HFD treatment

HFD has been previously observed to cause increase in body weight due
to accumulation of adipose tissue. In order to confirm this observation, our mice
were monitored weekly and their body weights were measured. Our results show
that HFD fed mice were significantly bigger than mice on Normal chow (NC).
There was 18-20% higher increase in body weight in HFD group after 14 days of
HFD feeding (Figure 5A), indicating that HFD was working on these mice as
expected. Additionally, blood biochemical analysis was performed at the end of
14 day HFD treatment to evaluate its effect on plasma glucose, insulin and fatty
acids. Our results demonstrated no changes in plasma glucose (Figure 5B) and
insulin (Figure 5C) after 14-day HFD feeding in mice, however, we observed
changes in fatty acid levels at this time point. Since plasma NEFA levels rise
after fasting due to activation of lipolysis (by Lipoprotein lipases) in adipose
tissues and the length of fasting determines NEFA concentrations [83], we took
blood samples at both fed and overnight fasting state to understand the
29
variations in fatty acid levels in our model. Significant increase in plasma fatty
acids or NEFA (non-esterified fatty acids) levels was seen in 14 days’ HFD fed
mice under fasting conditions (16-18 hour fast) (Figure 5D), indicating that this
group of mice were experiencing hyperlipidemic conditions at this time point.

Figure 5- 14 day HFD feeding increased body weight and caused
hyperlipidemia in mice. A, B, C, D. Body weight, Fasting blood glucose, insulin,
and NEFA (non-esterified fatty acids) plasma levels in NC vs HFD fed mice. *p
value<0.05, ***p value <0.0005.

Even though no hyperglycemia or hyperinsulinemia was seen after 14-day
HFD feeding, Glucose tolerance test (GTT) indicated slower clearance of glucose
in HFD fed mice during the 120 mins of the testing period (Figure 6A). This
indicates early occurrence of glucose intolerance and hence β cell dysfunction in
these mice. Furthermore, as shown by Insulin Tolerance Test (ITT), insulin
resistance in peripheral tissues was also observed in the mice fed HFD for 14
30
days. The insulin injected in these mice after 4-6 hour fasting was unable to bring
down blood glucose to normal levels after 90 mins of the test duration (Figure
6B), clearly indicating loss of peripheral insulin sensitivity. Although the length of
HFD feeding that can cause the development of insulin resistance is debatable,
many studies support its appearance after 1 week of treatment. As per a study by
Turner et al, 2013, insulin resistance in mice after 1 week of HFD treatment was
due to hepatic and adipose tissue insulin resistance followed by skeletal muscles
after 3 weeks [84].

Figure 6- Glucose intolerance and insulin resistance is seen in mice as
early as 14 days on HFD; GTT and ITT of mice fed with NC or HFD for 14 days
(A, B) along with area under the curve (AUC). On the line graph, blue line
represents NC mice plasma glucose while the red line indicates HFD fed mice
plasma glucose values. n=3 each group. *P value<0.05, ***p value<0.0005

Since we observed hyperlipidemia in our mice model after 14 days of HFD
treatment, we wanted to then study the effect of this condition on β cell
proliferation. Therefore, to look at the effect of HFD on β cell proliferation,
pancreas isolated from mice after treatment were subjected to Ki67 staining. Ki67
is a marker of proliferation and is only seen in cells that are undergoing cell
31
division. Our results indicated significant rise in β cell proliferation with 14-day
treatment causing 3-fold increase in HFD groups as compared to the NC
controls. These results support previously published data [67, 77] where short
term HFD feeding significantly increased β cell proliferation (Figure 7A,B).
However, β cell mass did not change significantly at this early time point (data
not shown).

Figure 7. HFD significantly increased proliferation in β cells of mice fed for
14 days. A. Ki67 and insulin double staining on 14 days HFD treated mice
pancreas, indicating β cell proliferation. Arrows indicate Ki67 positive cells within
islets. B. Quantification of the Ki67 positive insulin positive cells per total number
of beta cells. NC n=3, HFD n=5. Ki67- green, Insulin - red, DAPI- blue; **p
value<0.005.

In order to investigate the molecular players that caused significant
increase in β cell proliferation, pancreatic tissues from the treated mice along
with controls were subjected to Immunohistochemical (IHC) and western blot
analysis. As β cell replication is the main mechanism for adult β cell regeneration
[10] and Cyclins play a major role in regulating cell cycle, we decided to look at
Cyclin D1 and D2 expression in the β cells of mice treated with HFD. Cyclin D3 is
32
also expressed in very low amounts in the β cells but its role in proliferation is still
unclear. Although both Cyclin D1 and D2 have been reported to play an
important role in β cell proliferation, results have suggested that Cyclin D1 is
essential for the early postnatal β cell replication while Cyclin D2 is critical for
adult β cell growth [31]. We evaluated the expression of these cyclins in β cells
after 14 days HFD where hyperlipidemia is observed without hyperglycemia. The
islets from this cohort did not show significant increase in either of the Cyclin D
proteins (Figure 8 A, B). Therefore, lipids might be regulating β cell proliferation
through other cell cycle regulators that need to be further analyzed. Additionally,
high glucose levels may be required for the hyper-proliferative effects of HFD,
along with hyperlipidemia.
Figure 8-Cyclin D1 and D2 levels do not change in mice after 14 day HFD
treatment in mice islets. A. IHC quantification of cells positive for Cyclin D1 or
D2 and insulin per total insulin positive cells, B. Western Blot from islets isolated
from mice after 14 day NC and HFD treatment. IHC quantification (n=3 per
group) and western blot analysis (n=4 per group).
33

III-2-2 Cyclin D2 levels significantly increased in β cells after 2 month HFD
treatment leading to increased proliferation

The metabolic profile of 2 month HFD fed mice indicated 60% increase in
the body weight in HFD mice as compared to NC controls (Figure 9A), indicating
higher accumulation of adipose tissue mass with longer HFD treatment (as
compared to 14 day HFD treatment). Additionally, blood plasma screening
indicated the development of hyperglycemia after 2 months’ HFD feeding (Figure
9B). Compensatory increase in plasma insulin levels was also observed in the
HFD fed group, indicating that these mice were also experiencing
hyperinsulinemia along with hyperglycemia (Figure 9C).

Figure 9- 2 month HFD treatment cause increased body weight,
hyperglycemia and hyperinsulinemia. A, B, C. Body weight, Fasting blood
glucose and insulin levels in NC vs HFD fed mice. Both hyperglycemia and
hyperinsulinemia together indicate insulin resistance in HFD fed mice. *p
value<0.05, ***p value <0.0005.

Co-existence of hyperglycemia and hyperinsulinemia suggests insulin
resistance and in order to test that we performed GTT and ITT on these mice.
Both GTT and ITT analysis indicated glucose intolerance and insulin resistance
after 2 months HFD feeding (Figure 10 A & B). Quantification of Area under the
34
curve (AUC) confirmed the significant difference. This indicates that 2 month
HFD fed mice had both decline in β cell function along with decreased ability of
peripheral organs to metabolize glucose, which ultimately led to hyperglycemia in
these mice.
Figure 10- Glucose intolerance and insulin resistance is observed in mice
after 2 month HFD treatment. GTT and ITT of 2 months (A, B) HFD treatment
along with area under the curve (AUC). On the line graph, blue line represents
NC mice plasma glucose while the red line indicates HFD fed mice plasma
glucose values. ***p value<0.0005.

Moreover, significant increase in fed plasma NEFA levels was seen the 2-
months HFD treatment group (Figure 11A). This increase in the levels of NEFA
under HFD fed conditions might indicate increased levels of fatty acids in
circulation due to diet. However, we did not observe changes in fasting levels of
NEFA in NC or HFD fed groups. One theory to explain why 2M HFD treatment
group did not have significant increase in NEFA levels under fasting conditions is
that this group experienced high levels of insulin in their fasting plasma. Insulin is
known to decrease the activity of lipases that release fatty acids from adipose
tissues, thus decreasing the circulating levels of fatty acids.
35
Plasma Triglyceride (TG) levels along with fatty acids are indicative of
elevated presence of lipids in the blood and has been linked to dyslipidemia
under insulin resistant conditions. Thus, TG levels were measured in both NC
and HFD groups under both overnight fasting and fed states. Our results
indicated significant increase in plasma TG levels after 2 months of HFD
treatment (Figure 11B). Together, significant increase of fatty acids along with
TG level indicate that mice fed with HFD for 2 months are exposed to
hyperlipidemic conditions.
Figure 11- High levels of plasma NEFA and triglyceride (TG) levels indicate
hyperlipidemia in mice treated with HFD for 2 months. A, B. Plasma fed
NEFA (non-esterified fatty acids) and fasting Triglyceride (TG) levels in NC vs
HFD fed mice. *p value<0.05, **p value <0.005.

Therefore, the blood metabolic profile of mice fed with HFD for 2 months
indicated that these mice were exposed to both hyperglycemic as well as
hyperlipidemic conditions. Our next step was to evaluate the effect of such
conditions on β cell proliferation. Our Ki67 staining, done on the pancreatic
section of 2 months NC vs HFD fed mice, indicated approximately 13-fold
36
increase (from 0.12% in NC versus 1.5% in HFD group) in proliferation in HFD
groups as compared to the NC controls (Figure 12). This data is supportive of
other studies in which significant induction in β cell proliferation was observed
after few weeks of HFD treatment [77].
Figure 12- HFD significantly increased proliferation in β cells of mice fed
for 2 months; A. Ki67 and insulin double staining on 2 month HFD treated mice
pancreas, indicating β cell proliferation. Arrows indicate Ki67 positive cells within
islets. B. Quantification on the right, NC n=5, HFD n=4. Ki67- green, Insulin - red,
DAPI- blue; **p value<0.005.

In order to identify the molecular cell cycle regulators that might be leading
to proliferation, we next checked the levels of Cyclin D1 and D2 in the islets of
mice fed with NC and HFD. We did not see much differences in Cyclin D1
expression in β cells of NC vs HFD fed mice, but Cyclin D2 levels appeared to be
up-regulated after HFD treatment. Our IHC results confirmed 2-fold higher
expression of Cyclin D2 in the β cells of 2 month HFD treated mice as compared
to NC controls (Figure 13A). Western blot from islets isolated after 2 months’
treatment with HFD also corroborated significant increase in Cyclin D2 in HFD
group as compared to NC (Figure 13B). These results are consistent with
37
previously published study by Stamateris et al 2013 that has shown that HFD
induces β cell proliferation via Cyclin D2 [67]. They also published another study
recently that indicated that Cyclin D2 was essential for glucose-induced β cell
proliferation to occur [72]. Since HFD caused hyperglycemia (Figure 9B) as well
as hyperlipidemia (Figure 11 A, B) after 2-month treatment, both glucose and
fatty acids might be responsible for increased proliferation and Cyclin D2
induction.

FIGURE 13- HFD feeding for 2 months significantly increased Cyclin D2
levels. Immunohistochemistry (IHC) and Western blot analysis was performed to
study the levels of Cyclin D1 and Cyclin D2 in mice fed with HFD. A. Right panel,
immunostaining of Cyclin D2 (green) in 2 M HFD treated mice pancreatic islets,
left panel IHC quantification; B. WB of 2M HFD treated mice with quantification.
*p value<0.05.

The effect of glucose on Cyclin D2 has been established previously [71,
72]. To specifically address the effect of fatty acids on β cell proliferation, we
treated INS-1 cells with 0.4mM Palmitic Acid (PA) in the presence of
physiological concentration of glucose (6mM) for 48 hours. Our results indicated
38
that Cyclin D2 (as well as Cyclin A) are both induced after PA treatment for 48
hours (Figure 14B). This observation suggests that fatty acids can induce cyclin
levels in β cells independent of glucose to cause mitogenic stimulation.
There are several signaling pathways that are responsible for β cell
proliferation including, PI3K/AKT, mTOR and MAPK/ERK pathways. PI3K/AKT is
one of the major signaling pathways downstream that has been observed to
contribute to β cell proliferation. Constitutively active AKT and AKT
overexpression have been reported to cause β cell proliferation and bigger islets
by regulating levels of Cyclin D1, Cyclin D2 and p21 [18, 19]. Additionally,
mTOR, an effector of this pathway, not only responds to growth factor regulation
but also responds directly to nutrients [85, 86]. Inhibition of mTOR pathway by
using rapamycin caused reduced β cell proliferation, reduced pancreatic islet
content and reduced islet size [28]. Moreover, mice genetic models have
indicated that deletion of S6K (downstream of mTOR) lead to smaller islets in
mice that become glucose intolerant [25]. These studies indicate the importance
of PI3K/AKT and mTOR pathway in regulating β cell replication. We, therefore,
evaluated the expression of proteins involved in the PI3K/AKT/mTOR signaling
pathway.
Our data analysis suggests that the activity of S6K, the target of mTOR is
induced by the treatment of PA as indicated by the increased phosphorylation of
S6K. Additionally, our results support that this increase of S6K phosphorylation
occurs at the absence of AKT activation. AKT is a crucial pathway that regulates
cellular proliferation. Therefore, we expected to see AKT activation after
39
treatment, though no HFD studies have shown AKT activation in islets under
proliferative state in β cells. However, 7 month HFD feeding has been previously
reported to downregulate AKT activation by enhancing PTEN levels in islets [87].
Supporting these results, our in vitro treatment of INS-1 cells showed that
phosphorylation of AKT is inhibited by PA treatment, indicating downregulation of
AKT activity. As AKT phosphorylation is not induced whereas S6K is activated by
PA treatment, these data imply that the mTOR pathway is involved in PA
signaling independent of AKT (Figure 14A). Consistent with the role of mTOR
signaling in PA induced stimulation of cyclins, rapamycin treatment in INS-1 cells
decreased the expression of p-S6K which was accompanied by downregulation
of Cyclin D2 and Cyclin A (Figure 14B). This result further confirmed the
involvement of mTOR signaling.
FIGURE 14. In vitro treatment of INS-1 cell line with Palmitic Acid (PA)
induces p-S6K. A. PA treatment decreases AKT but increases p-S6K levels, B.
Rapamycin treatment inhibits p-S6K along with Cyclin D2 and Cyclin A.

In order to acquire more supporting in vivo data looking at the changes in
various canonical pathways in the islet transcriptome after HFD feeding, we
performed RNA-seq analysis on mice islets fed with HFD vs NC controls. RNA
40
seq is an emerging technology that helps in the in-depth analysis of cell
transcriptomes and studies are been published that are looking into β cell
transcriptomes of diabetic vs healthy individuals and treated vs untreated
human/mice islets [88]. Our unbiased enrichment analysis (we used Ingenuity
pathway analysis- IPA) using fold change (FC 1.5) revealed that p70S6K and
mTOR are one of the top few pathways that are significantly upregulated in HFD
treated mice as compared to their NC counterparts (Figure 15). The data is
expressed as z-score which is a score provided based on literature in their
database and our data. Positive z-score indicates activation while negative score
indicates inhibition of the pathway. As shown in the figure, majority of genes
involved in the mTOR pathway are upregulated and hence it has a positive z-
score. This result firmly suggests the involvement of mTOR pathway in β cell
proliferation after HFD treatment in mice.

41
Figure 15. Ingenuity Pathway Analysis (IPA) results showing significant
enrichment of the mTOR pathway in HFD islets as compared to the NC
controls at FDR (False discovery rate) 0.05 and Fold change (FC) 1.5.

Additionally, to corroborate the effect of mTOR on HFD induced β cell
proliferation in vivo, we treated the mice fed HFD with Rapamycin and then
observed its effect on β cell proliferation. Mice were fed HFD for 5-6 weeks and
injected rapamycin over the last 8 days (0.3mg/Kg/mouse everyday) (Figure
16A). Our results indicate significant reduction in β cell proliferation in mice
injected with rapamycin as shown by BrDU staining (Figure 16B). These mice
also had significant increase in their blood glucose (Figure 16C), which is
supportive of previously published data that indicates that rapamycin treatment
causes decrease in islet insulin content and leads to hyperglycemia is mice [28].
This result supports our in vitro effect and confirms the involvement of mTOR
pathway in HFD mediated β cell proliferation.

42
Figure 16. Rapamycin treatment abolishes HFD-induced β cell proliferation.
A. Treatment schematic, B. Quantification of BrDU positive insulin positive cells
in islets of mice fed HFD with and without Rapamycin treatment, C. Fasting
glucose levels significantly increase in Rapamycin treated mice as compared to
control treated HFD mice and NC fed mice. N=6 for HFD+control group and n=7
for HFD+rapamycin treatment. ***p value < 0.0005.

MAPK/ERK is another important pathway that has been previously seen to
have mitogenic effect on cells [89]. However, its role in β cell proliferation
induction is still unclear. Nevertheless, our treatment results with 0.4mM PA in
INS-1 cells consistently showed significantly increased phosphorylation of ERK
(Figure 17A), suggesting its activation and potential involvement. But when we
used MEK inhibitors (that inhibit phosphorylation of ERK through MEK) PD98059
and U0126, to identify if this kinase is required for the fatty acid mediated effect,
the decrease in p-ERK levels did not correlate with downstream Cyclin D2 levels
(Figure 17B). These results suggested that while ERK is a target for PA, it is not
responsible for the pro-proliferation effects we see in β cells. Role of ERK
activation by fatty acid stimulation needs to be further analyzed.

FIGURE 17. In vitro
treatment of INS-1
cell line with
Palmitic Acid (PA)
induces p-ERK but
does not mediate
Cyclin levels via
ERK pathway. A. PA
induces
phosphorylation of
ERK, B. MEK
inhibitors (PD98059
and U0126) inhibit ERK phosphorylation but do not downregulate Cyclin D2 and
Cyclin A.
43

III-3 Discussion-

Our results illustrate that HFD induces proliferation in mice after short term
feeding by increasing Cyclin D2 expression in β cells. In vitro data is supportive
and further demonstrates that fatty acids such as palmitic acid, in the presence of
physiological levels of glucose, is capable of inducing β cell proliferation. We
further demonstrated that palmitic acid signals the induction of β cell replication
through cell cycle regulatory proteins such as cyclin D2 via induction of mTOR
expression and mediated S6K induction. This novel finding is supported by our
results that indicate upregulation of Cyclin D2 expression upon palmitate
treatment, phosphorylation and activation of S6K which is downstream of mTOR
pathway and reversal of Cyclin D2 induction after rapamycin treatment that
inhibits mTOR. This finding for the first time clearly illustrated the consequence
of regulatory pathways stimulated by lipids to augment β cell proliferation.
β cells, located in islets of Langerhans, are the pancreatic cells that
release insulin and are responsible for maintaining glucose homeostasis.
Although adult β cells are mostly quiescent, they possess plasticity to proliferate
in order to meet increasing requirements under conditions such as pregnancy,
obesity and insulin resistance. β cell mass is maintained by a delicate balance
between β cell growth by replication & differentiation and β cell death and
dysregulation in β cell maintenance lead to diseases such as Diabetes.
Replication of preexisting β cells have been accepted to be the main mechanism
that regulates β cell mass. β cells respond to mitogenic stimuli such as nutrients
and growth factors by stimulating signaling pathways that can activate
44
downstream cell cycle proteins such as Cyclin Ds. Glucose is one such stimuli
that has been seen to activate β cell proliferation by up-regulating Cyclin D2 [72].
Since hyperglycemia and hyperlipidemia are both hallmarks of diabetes and
compensatory increase in β cell proliferation is seen during early/acute exposure,
we wanted to study how high glucose and lipids impact the various canonical
signaling pathways and enhance β cell mass.
We used a HFD model in which mice were fed HFD for short-term (14
days and 2 months) and then looked at the effects of the resulting hyperglycemia
and hyperlipidemia at β cell proliferation. In our study, we observed that 14 days
and 2 months HFD treatment significantly increased the number of Ki67 positive
β cells indicating augmented proliferation in these cells. Insulin resistance was
observed early on with 14 day HFD fed group showing decreased ability to
process insulin and decrease blood glucose. Hyperglycemia and
hyperinsulinemia was seen after 2 month HFD treatment, following insulin
resistance. This group also experienced increased levels of fatty acids and
triglycerides in their plasma at the end of study indicating higher exposure of β
cells to lipids.
When we looked at the cell cycle proteins that might be causing an
increase in β cell proliferation, we found that Cyclin D2 was significantly up-
regulated in mice that were fed HFD for 2 months. This result is similar to the
published work by Stamateris et al, that indicated upregulation of Cyclin D2 after
7 day HFD treatment in mice [67]. Even though we saw 3-fold increase in
proliferating cells in 14 day HFD fed mice, no upregulation of cyclin Ds was
45
observed. This may be due to absence of hyperglycemia in these mice, as
glucose has been established to be a critical modulator of Cyclin D2 expression.
However, these mice do experience hyperlipidemia which alone is unable to
upregulate cyclins (D1, D2 and A) in vivo, but might be inducing proliferation
through other regulatory proteins.
Since it is harder to separate the glucose and fatty acid effects in vivo, we
decided to employ an in vitro system wherein we treated INS-1 cells with 0.4mM
PA under physiological glucose concentrations (6mM). After treatment, we
screened for several pathways that might get induced and cause proliferative
effects in β cells including PI3K/AKT, mTOR and MAPK/ERK pathways. Our
results indicated decrease in levels of p-AKT, suggesting that this pathway does
not take part in fatty acid mediated cell division under our treatment conditions.
Several studies have been published looking into effect of palmitic acid on AKT
but they show varied results. While some show that PA causes AKT activation
[79], others claim that PA reduces active AKT and induces apoptosis [80, 81].
Although there was decreased activity of AKT after PA treatment, we observed
increased phosphorylation of S6K, a downstream target of mTOR pathway,
indicating AKT independent stimulation of mTOR pathway. Furthermore,
inhibition of mTOR pathway using rapamycin decreased Cyclin D2 levels,
indicating that this pathway may be involved in lipid mediated β cell proliferation.
mTOR has been previously shown to be activated by glucose and cause β cell
proliferation by upregulating Cyclin D2 [72], but its PA mediated activation is a
novel finding. Together, our results show that fatty acids such as palmitic acid
46
might be regulating mTOR expression and causing β cells to divide. This is
novel, as so far, the published studies have not been able to clearly demonstrate
pathways stimulated by lipids to enhance β cell proliferation. Furthermore, our
RNA seq results demonstrated that HFD treatment can significantly enrich genes
for mTOR pathway in the islets, suggesting that the pro-proliferative effect of
HFD might mediated through this pathway.
Future study will focus on understanding how fatty acids affect mTOR
pathway-directly or indirectly. One of the potential proteins that induces mTOR
activation when a cell is treated with fatty acids is PKC zeta. PKC zeta has been
recently shown by Lakshmipathi et al to be a master regulator of compensatory β
cell proliferation whose expression controls the activity of mTOR and
downstream Cyclin D2 during insulin resistant conditions [90]. PKC zeta is a
serine/threonine kinase which is activated by PI3K/PDK1 and has been reported
to be involved in cell replication, function, motility and survival [91]. GWAS
(Genome wide Association studies) have also associated several single
nucleotide polymorphisms (SNPs) to PRKZ gene (encodes PKC zeta) that have
been linked to increased risk of T2D development [92, 93]. All these above links
of PKC zeta to T2D and compensatory β cell proliferation makes it a very
attractive study target.
Overall, our study shows great potential by indicating the molecular
pathways stimulated by glucose and fatty acids in order to impact β cell
replication under insulin resistant conditions. Such knowledge can be used to
design targeted therapies for T2D patients who have decreased β cell mass and
47
their body is incapable of making enough insulin to maintain glucose
homeostasis.

48
Chapter IV: Long term High Fat Diet (HFD) decreases proliferation in β cells
by down-regulating CyclinD2 expression and up-regulating p27

IV-1 Introduction & Rationale-

Western lifestyle, including poor eating habits and sedentary
regime, have increased the incidence of diseases such as obesity. National
Diabetes Statistics Report has indicated obesity to be a leading cause of
Diabetes. Although pancreatic β cells are able to compensate for the increasing
demand under conditions such as obesity and insulin resistance, prolonged
hyperglycemia and hyperlipidemia proves to be fatal for β cells and causes
decrease in β cell mass and function. Long term exposure of β cells to glucose
(also described as glucotoxicity) have been observed to downregulate
expression of insulin along with various transcription factors such as Pdx1,
BETA2/NeuroD and MafA [94, 95]. Moreover, increased metabolic flux in
mitochondria causes excess production of Reactive oxygen species (ROS)
leading to oxidative stress. Oxidative stress has also been linked to decreased
insulin gene expression via JNK pathway [96]. Additionally, high glucose
environment causes the development of ER stress in β cells due to induction of
UPR and involvement of CHOP that ultimately leads to apoptosis [96].
Collectively, these studies clearly point to the detrimental effects of chronic
hyperglycemia in β cells. “Lipotoxicity” has been crafted as a term that describes
these detrimental effects of fatty acids on β cell dysregulation. Literature
published in the past few years have suggested that long term exposure to fatty
acids is detrimental to β cell function including decreases in glucose stimulated
49
insulin release (GSIS) as well as impaired insulin gene transcription in islets in
vitro [54].
Maintenance of β cell mass is as important as maintaining its function in
Diabetes. Islet mass is regulated by a fine balance between β cell proliferation
and cell death. There is accumulating evidence suggesting that long term
exposure of β cells to glucose and fatty acids causes increased cell death due to
apoptosis. A study in diabetic rats has indicated that these rats have decreased β
cell mass due to increased rate of apoptosis [55]. Diabetic rats also show upto 3-
7-fold increased in DNA laddering (indicative of apoptosis) as compared to lean
controls [97]. A 12 months High fat diet study in mice demonstrated decrease in
β cell proliferation and significant induction of apoptosis in obese, diabetic high
fat diet fed (HFD) mice versus controls [68]. Together these studies indicate that
β cell mass declines under diabetic conditions mainly due to induction of
apoptosis. In addition, rat islets when treated with high levels of glucose or fatty
acids independently in vitro showed higher levels of apoptosis which was
accompanied by increased expression of caspase 3 & bax (pro-apoptotic gene)
and reduced expression of bcl-2 (anti-apoptotic gene) [98]. This study illustrates
that its indeed the exposure of β cells to hyperglycemic and hyperlipidemic
conditions that leads to increased apoptosis and β cell death. Furthermore,
similar results were seen in human islets, where treatment of these islets ex vivo
with FFAs (2:1 ratio of oleate and palmitate) significantly decreased insulin
secretion and increased β cell death by activating caspases and regulating Bcl2
expression [99].
50
While proliferation is stimulated by hyperglycemia and hyperlipidemia with
short or moderate exposure, chronic exposure may lead to reduced proliferation
due to development of cellular senescence. A long term HFD study done by
Sone et al showed that after 12 months of high fat feeding the mice β cells have
decreased proliferation, increased apoptosis and upregulation of senescence
markers [68]. This study pointed out the role of senescence in diabetes and its
contribution to decreased β cell mass, as seen in Diabetic patients. p16
INK4a
and
p27
Kip1
, members of cyclin dependent kinase inhibitors (CDKIs) belonging to
INK4 and KIP family respectively, are very well known markers of β cell
senescence that are seen to be upregulated in aged animals [100]. p16
INK4a

expression increases with age and is negatively correlated with β cell
proliferation. Studies have indicated that while overexpression of p16
INK4a
in
young islets decreases proliferation potential, its deletion rescues the age-
dependent decrease in β cell proliferation [33]. Although no clear links of p16
INK4a
been made to Diabetes, a study performed by Pascoe et al demonstrated that
FFAs decrease glucose mediated β cell proliferation by induction of p16
INK4a
and
p18
INK4c
, in a 4-day glucose/lipid infusion mice model [56]. Together these
studies suggest the crucial role of p16
INK4a
in β cell proliferation. Additionally,
p27
Kip1
has been seen to be upregulated in β cells in genetic models of type 2
Diabetes- Insulin receptor substrate2 deletion (Irs2-/-) and leptin receptor
deletion (Lepr-/-). The same study demonstrated that overexpression of p27
Kip1

causes severe diabetes while its deletion p27
Kip1
prevented hyperglycemia by
increasing islet mass and improving insulin levels [35]. These results established
51
that p27
Kip1
is an important regulator of β cell mass and contributes to diabetic
phenotype in mice models. Together, it has been established that p16
INK4a
and
p27
Kip1
are senescent markers in β cells that may contribute to decreased islet
mass and proliferative potential in diseases such as Diabetes, however, how
their expression might be regulated under hyperglycemic and hyperlipidemic
conditions is not clearly known.
Even though the above literature implies the detrimental effect of long-
term exposure of glucose and FFAs on β cells, most of the models of chronic
exposure used in the studies are either lipid/glucose infusion, ex vivo treatment
with fatty acids or genetic obesity models which are not physiological and have
little clinical relevance. Therefore, we decided to use long term HFD feeding as
our experimental approach. HFD is a very well established model of obesity
induced Diabetes. Feeding mice long term with HFD will mimic exposure of
humans to high fat western fast food for prolonged periods and will help us to
study the effect of such long exposure of β cells to high levels of glucose and
fatty acids.
The goal of this aim is to investigate the effect of prolonged exposure of
high glucose and lipids/free fatty acids on β cell proliferation & mass and study
the involvement of cell cycle regulators such as Cyclins and CDKIs that might be
contributing to the effect.

52
IV-2 Results-

IV-2-1 β cell proliferation decreases after 12-14 month HFD treatment

To mimic chronic exposure to hyperglycemia and hyperlipidemia
condition, we used a physiological model by feeding 3 months old mice HFD
containing 60% fat in calories for 9 months and 12-14 months. At the end of the
study, we isolated pancreas from these mice and evaluated islet proliferation and
then elucidated the subsequent effect on islet mass. Our results showed that
after 9 months HFD feeding, there was no significant difference in proliferation
between NC and HFD groups (Figure 18A). We and others have previously
shown that short term exposure of β cells to HFD (upto 2 months) causes
increase in β cell proliferation as shown by Ki67 staining (Figure 12). Therefore,
no difference in proliferation between the NC and HFD groups after 9 months
HFD feeding indicates loss of β cell’s ability to respond to mitogenic stimulation
after chronic exposure to hyperlipidemia and hyperglycemia. Moreover, after 12-
14 months feeding, there was a significant reduction in proliferation of β cells
(Figure 18B), suggesting that the prolonged exposure of high glucose and fat
indeed reduces the capacity of β cells to regenerate.
The reduced β cell proliferation rate suggests that HFD might lead to
decreased islet mass, a phenomenon associated with Diabetes following obesity
and insulin resistance. We next quantified islet mass in 9 months’ and 12-14
months’ NC and HFD treatment groups. Surprisingly we observed significantly
higher islet mass in both HFD treated groups comparing to their respective
normal chow controls. We hypothesize that the accumulation of cellular divisions
53
over a period of time may have led to this effect. Indeed, lower islet mass is
observed in the 12-14 months vs. the 9 months HFD group, indicating that
exposure to HFD longer than 14 months may results in decrease in islet mass
compared to a matched normal chow control. The fold increase in islet mass in
HFD fed group vs normal chow was 2.9-fold compared to the 3.5 fold observed in
the 9-month group, indicating comparative decline in mass due to
glucolipotoxicity (Figure 18C).

FIGURE 18- HFD for 9-14 months decreases β cell proliferative potential
and subsequent islet area decrease. A. Ki67/BrDU and insulin double staining
on 9month and 12-14 month HFD treated mice pancreas, indicating decrease in
β cell proliferation. Arrows indicate Ki67/BrDU positive cells within islets.
Quantification on the right; B. Quantification of islet area measured as ratio of
islet area by total pancreas area in H& E stained slides in both 9M and 12-14 M
NC & HFD treatment groups. Ki67/BrDU- green, Insulin - red, DAPI- blue; *P
value<0.05, ***p value<0.0005.

β cell mass is regulated by the balance between β cell growth via
replication and β cell loss through apoptosis, necrosis and senescence. We also
evaluated the rate of apoptosis in the β cells of 12-14 months’ HFD fed mice.
54
TUNEL staining is a common method to detect DNA fragmentation in cells, which
is a hallmark of apoptosis. We could not detect any TUNEL positive β cells in the
pancreatic islets in either groups of mice. Detection of apoptosis event has been
difficult in β cells when not stimulated due to the low activity apoptotic levels.
To establish a correlation between the islet mass, β cell regeneration and
the metabolic changes induced by HFD, we determined fasting glucose and
insulin levels. Although body weight significantly increased in all the HFD fed
groups (Figure 19A), some loss of body weight was seen in 12-14 HFD mice
towards the end of the study. This is consistent with a previously published long
term HFD feeding study [68]. Hyperglycemia was observed in the 9 months’ HFD
group, while the 12-14 months’ mice did not show any significant difference in
their glucose levels (Figure 19B). The mechanism of this absence of
hyperglycemia in 12-14 months’ HFD fed mice compared to NC control is
unclear. Plasma insulin, on the other hand, was significantly higher in both 9
months’ and 12-14 months’ HGF groups (Figure 19C). But, the fold increase of
insulin dropped down 4 times in 12-14 months HFD mice. This observation may
be due to lack of hyperglycemia or reduced glycemic load in these mice and
hence reduced requirement of insulin secretion to maintain homeostasis.
55

FIGURE 19- Blood Biochemical analysis of mice treated with HFD for 9
months and 12-14 months. A, B, C. Body weight, Fasting blood glucose, insulin
levels in NC vs HFD fed mice. *p value<0.05, **p value <0.005, ***p value
<0.0005

Since glucose intolerance and insulin resistance are commonly observed
in diabetic patients, we decided to study these in our mice models using
intraperitoneal GTT (Glucose Tolerance Test) and ITT (Insulin Tolerance Test).
GTT in both groups showed poor clearance of glucose as seen by bigger area
under the curve for HFD group, indicating β cell dysfunction while ITT indicated
poor sensitivity of the peripheral organs to insulin. Quantification of area under
the curve (AUC) shows significant difference between NC and HFD groups
(Figure 20). Both 9 months’ and 12-14 months’ HFD treated mice experienced
56
hyperinsulinemia, which together with peripheral insensitivity indicates severe
insulin resistance in these mice.

FIGURE 20- Glucose intolerance and insulin resistance after 9M and 14M of
HFD feeding; GTT and ITT of 9M (A, B) and 14M (C, D) HFD treatment along
with area under the curve (AUC). Blue line indicates NC and orange line
indicates HFD mice average plasma glucose (mg/dL). *P value<0.05, **p
value<0.005.

IV-2-2 Cyclin D2 levels significantly decrease while p27 levels increase after long
term HFD treatment

Since there was reduction in β cell proliferation after 14 months HFD
treatment, we evaluated the levels of various Cyclin expression in these mice.
Cell cycle is regulated by Cyclins such as Cyclin Ds, A, B and E. As we either
could not detect or detect no difference in Cyclin B& E and A respectively in the
islets of the HFD and NC fed mice, we decided to just focus on Cyclin D1 and
D2. Cyclin D1 and D2 have been reported to play an important role in β cell
proliferation [31]. Immunohistochemistry of pancreatic sections from 9M HFD
57
treated mice indicated decreased levels of Cyclin D2 in β cells of these mice as
compared to NC controls (Figure 21A). These results were consistent with the
lower CyclinD2 levels in isolated islets from the 14M HFD treated mice, as seen
in the western blot (Figure 21B). However, Cyclin D1 levels did not change much
between NC and HFD fed groups after 9M (data not shown) and 14M treatment.
These results indicate that Cyclin D2 levels are negatively impacted by prolonged
exposure of glucose and fatty acids/lipids and may have been the cause of
decreased β cell proliferation.
FIGURE 21- HFD feeding for 9M and 14M significantly decreased Cyclin D2
levels. A. Immunohistochemistry (IHC) was performed to study the levels of
Cyclin D2 in mice fed with HFD for 9 months. Immunostaining of Cyclin D2
(green), Insulin (red) and DAPI (blue) in mice pancreatic islets, Right panel,
Quantification of IHC. B. Western Blot from islets of 14M HFD treated mice; p
value<0.05.

As cell cycle inhibitors such as p16 and p27 have been observed to
accumulate in the nucleus of β cells in diabetic models and cause decrease in
proliferation potential, we evaluated the levels of p16 and p27 in the islets to
check the effect of HFD on these proteins. Our results indicated no significant
58
difference in p16 expression levels among 9M NC & HFD mice groups and it was
not detected in the islets of 12-14 months HFD/NC treatment mice (data not
shown). This implies that p16 levels might not be regulated by hyperglycemic or
hyperlipidemic conditions. However, our western blot analysis confirmed the
increase in p27 levels in the islets of 12-14 months HFD treated mice (Figure
5A). Studies have shown that p27 is a senescent marker that accumulates in
terminally differentiated β cells [34] and its overexpression in β cells causes
severe diabetes [35]. Our results support accumulation of p27 in islets in mice
fed with HFD for 12-14 months. Thus, implying that p27 might be regulating the
decreased proliferative potential of β cells under long term exposure of high fat
diet conditions.
Together our data indicates, decrease in Cyclin D2 and increase in p27
might be responsible for decreased β cell proliferation in long term HFD fed mice.

IV-2-3 Chronic high levels of Triglycerides (TG) may be responsible for increased
p27 and decrease in β cell proliferation.
Hyperlipidemia is yet another common symptom in diabetes and hence we
decided to analyze the levels of Non-esterified fatty acids (NEFA) and
triglycerides (TG) in all groups. We analyzed both fasting and fed states as fatty
acids are released under fasting conditions and their concentration varies with
the length of fasting. Our results indicate non-significant differences between NC
vs HFD groups in both 9 months’ and 12-14 months’ treatment conditions under
both fasting and fed states (Figure 22A). However, we observed a significant
59
increase in plasma Triglyceride (TG) levels in fed plasma of 12-14 months HFD
group mice (Figure 22B). One theory that could explain such an observation is
that since plasma NEFA is the main substrate for hepatic TG production (in the
form of VLDL-TG), higher levels of NEFA due to HFD are causing more VLDL-
TG secretion and hence higher plasma TG concentrations [101]. Although
several studies have shown beneficial effects of cellular TG accumulation in β
cells [102, 103], effect of plasma TG on β cells is still debatable. Our data
indicated a negative correlation between plasma TG and islet mass, and β cell
proliferative potential. Thus, we hypothesized that this elevated TG may serve
as a signal that causes decreased β cell mass and proliferation in these mice.

FIGURE 22- Triglyceride (TG) levels significantly increase in HFD fed mice
as compared to NC control mice; A, B. Plasma NEFA and TG levels under
fasting and fed state. *p value<0.05.

There is no literature in our knowledge so far that indicates or links β cell
proliferation with plasma TG levels. Our analysis here observed that elevated TG
levels together with decreased β cell proliferation (Figure 23A) and
downregulation of Cyclin D2 levels. We also observed increased expression of
60
p27 in islets of HFD fed mice (Figure 23A). To address the specific effect of TG
on β cells, we used the INS-1 (rat insulinoma) cell line. We treated INS-1 cells
with 30, 60 and 120ng/ml TG for 24 and 48 hours and investigated the
expression of Cyclin D2 and p27. Our results showed that 24 and 48-hour
treatment of cells with TG had a dose dependent effect on Cyclin D2 thereby
downregulating its expression (Figure 23B). Whereas, all the treatment doses of
TG and time points caused upregulation of p27 (Figure 23B). 48-hour treatment
with 120ng/ml TG concentration was too toxic for cells. This indicates that high
plasma TG levels due to long term HFD treatment may be a contributing factor
that decreases β cell proliferation and causes β cell decline, likely by inducing β
cell senescence.

Figure 23- Increased Triglyceride levels cause accumulation of p27 in β
cells after 14M HFD treatment causing decrease in proliferative potential. A.
In vivo data- Graph summarizing fold change in proliferation among HFD vs NC
groups after 9M and 14M HFD feeding; Plasma TG levels increase significantly
61
in 12-14M HFD group; Increased expression of p27 as seen in western blot in
14M HFD islets as compared to NC controls, B. In vitro treatment of TG in INS-1
cells causes dose dependent increase in p27 levels after 24 and 48 hrs. *p
value< 0.05

IV-2-4 Treatment with saturated fatty acids in vitro causes decreased viability,
proliferation and increased apoptosis.
In obese individuals, there is increase in adipose tissue mass, as
compared to lean controls, which stores as well as releases large quantities of
fatty acids. These fatty acids have been reported to be responsible for insulin
resistance in the peripheral organs. They have also been observed to have both
pro and anti-proliferative effects on β cells. Although there are literatures that
supports both sides of the argument, no clear conclusion can be made on its
effect on β cell proliferation. In our long term HFD treatment model, we observed
increased insulin resistance and decreased β cell proliferation, accompanied by
increase plasma triglyceride levels. However, due to the low abundance of cells
stainable for TUNEL, we were unable to address the toxic effect of chronic lipid
exposure to HFD on β cells using the in vivo model. Therefore, using the INS-1 β
cells, we investigated the effect of palmitic acid (PA) (which is the major
saturated fatty acid in our diet) treatment on β cell viability.
In order to assess the viability of INS-1 cells when treated with different
concentrations of PA, MTT assay was performed. MTT assay is a colorimetric
method to determine cell metabolic activity. Cells were incubated with PA for 24
and 48 hours followed by addition of tetrazolium dye MTT 3-(4,5-dimethylthiazol-
2-yl)-2,5-diphenyltetrazolium bromide. As shown, the results indicated dose
62
dependent decrease in cell viability at both 24 and 48 hours’ time points (Figure
24A). We chose 0.4mM concentration of PA for future experiments since this
concentration caused intermediate effects on cells (30-40% loss). We also
performed cell count assay to determine the effect of PA on β cell growth/death.
Our results indicated that, PA treated cells had lower cell count as compared to
the BSA treated control cells (Figure 24B). BSA was used as a carrier to
solubilize PA for the in vitro treatments and hence was used as a negative
control. These results indicate that PA has a negative impact on growth.
Since PA influenced β cell growth, for in-depth investigation of the cell
cycle we used Cell Cycle FACS (Flow cytometry) analysis on PA treated cells.
Cell cycle analysis utilizes dyes such as Propidium Iodide to stain the DNA of
permeablized cells such that the fluorescent intensity correlates directly with the
amount of DNA. After evaluating all phases of cell cycle i.e. G0, G1, S and G2
phase in the treated cells, our results show that PA treatment has an impact on
the S phase. Although the percentage of cells in S phase varied with each set of
runs, each set showed reduction in S phase when treated with PA as compared
to the no-treatment control (Figure 24C). Cells were treated with FBS as a
positive control to confirm significant increase in cell division capability in
presence of serum and hence it showed more cells in the S phase.
As previous studies have indicated the effect of FFA such as PA on β cell
apoptosis and death, we performed Annexin V/ Propidium Iodide (PI) FACS
analysis on INS-1 cells to interrogate the effect of PA on β cell death. Cells
undergoing apoptosis translocate phosphatidylserine (PS) from the inner side of
63
the membrane to the outside, which can then be labeled by Annexin V. On the
other hand, PI stains dead and damaged cells. Hence, both Annexin V and PI
together, stain for early and late apoptotic/dead cells in the pool. Our data
supports the increase in the number of apoptotic cells (both Annexin and PI
positive) after PA treatment as compared to both no treatment control and BSA
treated cells (Figure 24D). Additionally, when we looked at the molecular
mechanism leading to apoptosis and cell death we found that PA treated cells
have higher expression of cleaved caspase 3, an executioner caspase that plays
a central role in cell apoptosis.
Together these results clearly indicate that saturated fatty acid (PA)
treatment causes decrease in β cell viability and growth (S-phase) along with
increase in cell apoptosis (as shown by both FACS and western blot).

Figure 24- Palmitic Acid (PA) decreases cell viability, growth and increases
cell apoptosis in vitro. A. Bar graphs showing MTT assay performed in INS-1
64
cells using different concentrations for PA for 24 and 48 hours, B. Growth curve
done using 0.4mM PA and 1% BSA (as control) for 8 days, C. Cell cycle FACS
analysis. Bar graph showing the percentage of cells in S-phase after treatment
with PA; Without serum and with serum being negative and positive controls, D.
Annexin V/Propidium Iodide FACS data with quantification, Western blot showing
cleaved caspase 3 expression in PA treated cells. *P value<0.05, **p
value<0.005.

IV-3 Discussion

In this long term HFD study we observed significant decrease in β cell
proliferation along with down-regulation of Cyclin D2 and up-regulation of p27 in
pancreatic β cells after 14 months of HFD treatment. This was accompanied by
hypertriglyceridemia in the HFD fed mice group. Our in vitro treatment of INS-1
cells with TG caused reduction of Cyclin D2 along with induction of p27,
supporting our hypothesis that hypertriglyceridemia may be responsible for
decreased β cell proliferation. So far in our knowledge, this is the first study that
has shown the effect of triglycerides on cell cycle modulators and its
consequence in decreased proliferation in β cells.
We used a long term HFD model in order to mimic hyperglycemia and
hyperlipidemia seen in Diabetic individuals. Our 9-14 months HFD feeding model
was aimed at figuring out the molecular mechanisms that affect β cell
proliferation and mass due to prolonged high levels of glucose and lipids/fatty
acids. HFD has been seen to affect β cell proliferation and eventually mass,
depending on the length of exposure. Two studies have been published so far
with long term (1 year) HFD treatment in mice. Study by Hull et al 2005,
observed increase in β cell mass due to hyperplasia along with impaired β cell
function after 1 year HFD feeding [69]. Additionally, Sone et al 2004 reported
65
decrease in β cell proliferation, increase in apoptosis and induction of
senescence at the end of a yearlong HFD feeding [68]. But both these studies
have been unable to clearly identify the molecular players that are affected by
long term exposure of glucose and lipids which can affect proliferation or death in
β cells.
Usually β cells have a very slow turn-over rate in adults but in mice they
have been observed to proliferate to adapt to various insulin resistant conditions
such as pregnancy and obesity. We and others have observed significant
increase in β cell proliferation when mice were fed HFD for 2-8 weeks’ indicative
of the ability of β cells to respond to mitogenic and metabolic stimuli. However, in
our study, 14 months’ HFD treatment in mice caused significant decrease in β
cell proliferation as compared to NC controls. This was accompanied by down-
regulation of Cyclin D2 and up-regulation of p27 in β cells of HFD fed mice β
cells as indicated by the western blots and IHC data. Cyclin D2 is an essential
cyclin involved in G1/S phase of the cell cycle. Several studies have shown its
contribution to β cell proliferation in response to mitogenic stimuli such as
glucose, wherein glucose stimulation increased β cell proliferation by regulating
Cyclin D2 levels [71, 72]. These studies and our results suggest that Cyclin D2
may act as an “adaptive molecule” contributing to the adaptive ability of β cells to
increase proliferation when required, but decreases when the cell undergoes
replicative senescence. P27, on the other hand, is a CDK inhibitor that restricts β
cells from going into the cell cycle division phase. Its higher expression in the
islets after long term HFD treatment is suggestive of induction of senescence in β
66
cells. Previously, deletion of p27 in genetically induced diabetic models (Irs2
-/-

and Lepr
-/-
) was reported to reverse diabetic phenotype and cause compensatory
β cell proliferation [35]. However, senescence is not very well understood in β
cells. One of the long term HFD studies suggested that β cells undergo ROS
mediated senescence due to induction of p38 MAPK pathway which leads to
elevation of p21 expression [68]. P21 belongs to CDK inhibitor CIP family and
has been previously reported to be induced by oxidative stress in rat islets
treated with H
2
O
2
and in islets of Zucker diabetic fatty rats [104]. However, we
were unable to detect p21 in our NC and HFD fed mice islets. Our study
suggests that decrease in Cyclin D2 and increase in p27 expression might cause
the β cells to undergo replicative senescence leading to decreased β cell
proliferation after chronic exposure to high glucose and fat in mice.
Contrary to our expectations, the levels of glucose and fatty acids were
similar in NC and HFD fed groups after 14 months’ HFD treatment. We did not
observe hyperglycemia or hyperlipidemia in HFD fed group. Therefore, we could
not relate the effect on β cell proliferation to glucotoxicity or lipotoxicity. However,
levels of plasma triglycerides were significantly higher in the HFD group. In order
to interrogate the effect of TG on β cells, we used INS-1 cells and treated them
with TG. The results confirmed that TG can have a negative impact on
proliferation by downregulating Cyclin D2 levels while increasing the expression
of p27. In pancreas, circulating TG are broken down into fatty acids by islet
lipases and these fatty acids are then taken up by the β cells either by passive
67
diffusion or through fatty acid receptors such as CD36 [105]. Hence, effect of TG
in vitro can be indirectly related to effect of fatty acids in vivo.
Further research is required to confirm the mechanism by which lipids
directly or indirectly regulate β cell proliferation. It is possible that proteins
affected by glucose, may also respond to or are affected by fatty acids as both
are major nutrient sources for β cells. Menin, product of MEN1 tumor suppressor
gene, has been previously reported to negatively regulate β cell proliferation in
mouse models of obesity and pregnancy. At the molecular level, MEN1 has been
observed to regulate transcriptional expression and upregulation of p21 and p27
[106]. Furthermore, it has also been published that glucose can downregulate
Menin through microRNAs (miR-17) and promote proliferation in β cells [107].
Together, these studies make it plausible that Menin might be a molecular
regulator that is driving the expression of p27 under chronic hyperlipidemic
conditions.
In summary, our results show that long term exposure to fatty acids/TG
causes decrease in β cell proliferation by affecting the cell cycle regulatory
proteins. Both in vivo and in vitro data support the involvement of Cyclin D2 and
p27, which show potential to be used as targets for β cell regeneration therapy. A
better understanding of the molecular modulators that contribute to the
decreased β cell mass under diabetic conditions will help in improving β cell
number in patients experiencing severe symptoms and hopefully in future can
replace insulin therapy.

68
Chapter V: Overall Discussion

Our study confirmed that 1) short term HFD induces proliferation and
chronic exposure to HFD leads to the inability of β cells to compensate; it
supported that 2) High plasma levels of glucose and fatty acids correlate with β
cell proliferation while high levels of triglycerides (TG) are associated with lack of
proliferative response; and for the first time demonstrated that 3) HFD induced
proliferation is dependent upon S6K and mTOR signal, independent of AKT. ERK
is induced by fatty acids but unlikely to contribute to lipid-induced β cell
proliferation, 4) Chronic exposure related loss of β cell compensation is related to
increases in plasma TG levels, and 5) TG induces senescence in INS-1 cells like
HFD does in vivo by induction of p27. These conclusions were supported by
increase in β cell proliferation, accompanied by increase in the levels of plasma
glucose and fatty acids (fed state) after 2 months’ HFD exposure. Cyclin D2 was
significantly upregulated at this time point which could explain the robust levels of
proliferation at this time point. RNA seq data from the islets indicated
upregulation of mTOR pathway genes in HFD treated islets as compared to NC
counterparts. Moreover, rapamycin treatment in mice abrogated the HFD
induced proliferation, supporting the involvement of mTOR pathway in β cell
proliferation. Additionally, our in vitro treatment of INS-1 cells with palmitic acid
(PA) indicated induction of mTOR pathway by fatty acids which caused
upregulation of Cyclin D2 in β cells. However, PA decreased levels of p-AKT
indicating that mTOR activation took place independent of AKT. Long term
feeding of HFD (12-14 months) in mice resulted in reduced proliferation rates,
69
associated with decreased cyclin D2 levels. These mice also showed higher
levels of p27 in their islets. Although these mice did not have high levels of
glucose or fatty acids in their plasma, their plasma triglyceride (TG) levels were
significantly higher. In vitro treatment of INS-1 cells with TG resulted in up-
regulation of p27 and down-regulation of Cyclin D2 expression levels indicating
that high levels of TG in plasma might be decreasing proliferation levels in mice
fed HFD for 12-14 months by affecting both p27 and Cyclin D2 levels. Since p27
is a marker of senescence, we could support that TG is causing β cell
senescence by inducing p27 levels both in mice islets and INS-1 cells.
Together our results validate previously published studies that indeed fatty
acids have an exposure time based effect on β cell proliferation, where short
term exposure is beneficial and causes increase in proliferation while long term
exposure is detrimental and results in significantly lower levels of β cell
replication. We also identified major players that regulate β cell proliferation
through lipid, in our study i.e. mTOR, Cyclin D2 and p27.
Our results support that Cyclin D2 plays an important role in the adaptive
proliferation of adult β cells when required under conditions such as insulin
resistance and obesity. Our HFD diet model depicts different stages of β cell
proliferation which are guided by the length of exposure to HFD. Increasing
levels of plasma glucose, fat and insulin resistance in peripheral tissues such as
muscle and adipose tissue, signal β cells to increase their mass and function,
which is translated into increase in levels of plasma insulin along with enhanced
β cell replication. During this phase, β cells express higher levels of Cyclin D2
70
that helps the cells to progress into the cell cycle. However, prolonged exposure
of β cells to the above-mentioned conditions reverses their tendency to
proliferate and they lose their ability to respond to mitogenic stimuli (as seen in 9
months and 14 months’ time points). This is correlated with the decreased
expression of cyclin D2 in β cells. Therefore, it is appropriate to call Cyclin D2 an
“Adaptive molecule” that helps β cells to adapt to different environmental cues by
aiding in expansion of β cell mass via replication of adult β cells.
Both glucose and fatty acids act as energy source for β cells. Glucose
increases ATP/ADP ratio in the cell which leads to closing of the K
+
ATP
channels, depolarization of the membrane, opening of the Ca
2+
channels and
finally insulin release. This process is referred to as GSIS (Glucose stimulated
insulin secretion). When plasma glucose concentrations are low, fatty acids help
in the insulin release and hence substantiate β cell function [76]. Glucose and
fatty acids independently have also been observed to increase cell cycle
progression in β cells. While glucose has been reported to activate insulin
pathway and downstream PI3K/AKT and mTOR signaling to stimulate β cell
replication [108], key molecular players modulating the effect of fatty acids still
remain mostly unknown. Our results indicated activation of mTORC1/S6K
pathway by fatty acids. mTORC1 activation has been previously implicated with
increased β cell size and proliferation. Deletion of ribosomal S6 kinase (S6K),
downstream translational regulator of mTORC1 pathway, leads to smaller mice
with smaller pancreatic islets and hypoinsulinemia [24, 25]. Additionally,
treatment with mTOR inhibitor, rapamycin, causes reduction in islet size and
71
proliferation [28]. On the other hand, β cell specific deletion of TSC2 (negative
regulator of mTOR pathway) leads to increased β cell proliferation, cell size and
hyperinsulinemia [26]. Recently glucose has been identified as a major mitogen
to induce mTOR via IRS2 in a mouse glucose-infusion model [72]. However,
activation of mTOR pathway by HFD has not been reported so far. Our RNA seq
data along with in vitro treatment of INS-1 cells indicated the up-regulation of
mTOR pathway under glucose/lipid treatment conditions. This is a novel finding
and would help to identify the role of glucose and fatty acids in β cell proliferation
in conditions such as obesity and diabetes. However, our research is still in initial
stages and needs further in vivo experiments to confirm.
Insulin resistance in organs such as muscle and adipose tissue precedes
diabetes development due to which these organs experience reduced sensitivity
to insulin, decreased glucose absorption and contribute to increase in plasma
glucose levels. Fatty acids released from adipose tissue have been reported to
induce insulin resistance by affecting insulin pathway [109]. Earlier studies have
shown that mTOR pathway might be responsible for induction of insulin
resistance in peripheral tissues such as skeletal muscle and adipose tissue by
downregulating insulin receptor substrate proteins with subsequent reduction in
AKT phosphorylation [110, 111]. However, no link between fatty acids and mTOR
pathway in the peripheral organs has been established. This would be an
interesting future study that would help in establishing the differential interaction
between fatty acids and mTOR in different organs (pancreas vs muscle &
adipose tissue) in obesity and would contribute to our current knowledge pool.
72
Additionally, hypertriglyceridemia is one of the hallmarks of diabetic
dyslipidemia and is linked with insulin resistance in obese and type 2 diabetic
subjects [112-114]. Elevated levels of triglycerides (TG) result from increased
production and decreased clearance of triglyceride-rich lipoproteins. A recently
published study by Liu et al 2016, studied the effect of TG on β cells independent
of other circulating factors such as glucose and insulin by using ApoC3
transgenic mice model, and showed that high levels of TG alone are not enough
to cause β cell dysfunction in upto 7 months old mice [115]. In our study, we
observed significantly higher levels of TG in mice plasma treated with HFD for
12-14 months, along with significant decrease in β cell proliferation and up-
regulation of p27 in islets. Together, the previously published data and our
results indicate that lipids/TG may not effect β cell mass or function in younger
mice (upto 7 months old) but has a negative impact on β cell proliferation in older
mice (15-17 months) and can cause β cell senescence by upregulating p27, a
senescence marker. P27 has been previously shown to accumulate in terminally
differentiated β cells during embryogenesis [34]. Senescence is the term that
describes replicative arrest in cells after a limited number of cell divisions.
Although senescence can occur through both telomere-dependent and
independent pathways, senescence could not be prevented in human primary
pancreatic β cells by expression of telomerase that extends the length of
telomeres [116] indicating major involvement of the telomere-independent
pathways. Additionally, many studies have reported ROS to cause cellular
senescence by induction of MAPK pathway such as extracellular signal-regulated
73
kinase (ERK), c-Jun N-terminal kinase (JNK) and p38 MAPK [117, 118]. In our
study, in vitro treatment of INS-1 cells with PA caused activation of ERK (as
indicated by upregulation of phosphor-ERK) but did not induce downstream
Cyclins, indicating its non-involvement in β cell proliferation. However, this
activation of ERK pathway may have a role in fatty acids induced oxidative stress
and ROS mediated cellular senescence in β cells in our system. But, more
markers such as p38 MAPK and p21 need to be studied in our system to support
the findings. In summary, our results support that lipids may cause senescence
in β cells by inducing markers such as p27 but the mechanism that causes
cellular senescence needs to be further evaluated.
Various treatment designs have been used to study the effect of
glucose/fatty acids on β cells and they have used methods such as infusion of
glucose/lipid emulsion into the mice for few days [56] or treating mice islets ex
vivo [49]. Though, these studies have the advantage of simpler experimental
design and reduced complication brought about by other tissues, they however,
lack physiological relevance and cannot be related to patients who are obese.
Therefore, we decided to address the role of glucose and fatty acids in an in vivo
model where hyperglycemia and hyperlipidemia is induced by high fat diet
feeding, mimicking human conditions of obesity and diabetes. In vivo studies,
however are challenged by interpretation due to crosstalk between pancreas,
liver, adipose tissue & muscle and the difficulty to pin point cause and effect
relationship. Moreover, diet induced obesity (DIO) and diabetes is very strain
dependent in mice. Few studies have shown that C57BL6 mice are more
74
susceptible to it than other strains and hence have been better characterized
[66]. Therefore, although we used a relevant mice model in our study, we are
aware of its limitations.
Together, our study provides a comprehensive look into the β cell
proliferation and mass changes that occur due to HFD feeding. In our knowledge,
this is the first study to evaluate longitudinal changes of β cells with both very
short (14 days) and very long (14 months) HFD exposure. This is also the first
study to show that HFD can influence mTOR signaling pathway to promote β cell
proliferation after short term exposure and induce p27 to restrict growth after long
term exposure. Maintaining β cell mass is important to avoid onset of diabetes. β
cell proliferation and mass changes according to the nutrients they are exposed
to. Both obesity and diabetes are associated with hyperlipidemia and hence
understanding the molecular regulation of β cell mass by fatty acids/lipids would
provide a better insight into development of new therapeutic targets for Diabetes.

75
Chapter VI: Future Directions

Results from our study indicated that mTOR signaling might be involved in
lipid mediated proliferation in β cells after short-term exposure to HFD. mTOR
pathway was also upregulated after treatment of INS-1 cells with Palmitic Acid
(PA) in vitro. Hence, our logical next steps are to study the relationship between
mTOR pathway and lipids and if lipids directly interact with mTOR to activate the
pathway.

VI-1 To determine if mTOR is a fatty acid sensor.
VI-1-1 Background & Rationale-
mTOR pathway responds to and integrates nutrient stimuli such as
glucose, amino acids and various growth factors, to control growth and
metabolism. Several studies have been able to outline the path followed by
amino acids to activate mTOR pathway. Amino acids are the building blocks of
proteins and are also involved in energy production by providing substrate to the
Krebs cycle. Early experiments indicated that deprivation of amino acids can lead
to autophagy in cells along with suppression of mTORC1 signaling [119],
suggesting a possible connection between amino acids, mTORC1 and
autophagy. Later it was revealed that amino acids interact with small GTPases
called Rags and activate them, by regulating nucleotide loading, so that they can
physically bind and activate mTORC1 downstream [120, 121]. However, Rags do
not directly activate mTORC1 [121]. Rags regulate the subcellular localization of
mTORC1 and direct its translocation onto the lysosomal surface through
76
Ragulator, complex of three Rag-binding proteins [122]. Translocation of
mTORC1 to the lysosomal surface brings it to close proximity of Rheb, which is
present on the lysosomal membranes, and this interaction between Rheb and
mTORC1 causes the activation of this pathway.
Glucose has also been observed to be responsible for activation of mTOR
pathway and cause induction of β cell proliferation. A study by Dickson et al,
reported the activation of mTOR signaling by glucose, independent of AKT (also
called PKB) in pancreatic INS-1 cells [123]. INS-1 cells in this study were treated
with or without IGF-1 in presence of either 3mM or 15mM glucose in a time
dependent manner. Results indicated activation of PKB by IGF-1 treatment with
no effect of glucose and p70 S6K (downstream of mTOR) upregulation by both
glucose and IGF-1. Inhibition of PKB signaling by treatment of Wortmannin,
partially reduced p70S6K levels, indicating that glucose can independently
activate this pathway. Additionally, a recent study by Stamateris et al investigated
the effect of glucose on insulin signaling pathways that can induce β cell
proliferation [72]. This study utilized a 7-day glucose-infusion mouse model and
demonstrated that glucose mediated β cell proliferation occurs through IRS2,
mTOR and Cyclin D2. Insulin receptor knockout or knockdown using shRNA,
however, did not have any effect on PCNA or BrDU levels in islets ex vivo,
indicating that insulin receptor is not required for glucose mediated β cell
proliferation. Together, these studies indicate that mTOR pathway responds to
glucose and mediates β cell proliferation.
77
However, role of mTOR in response to lipids remains unclear. Our results
indicate the involvement of mTOR pathway in a lipid dependent manner to
promote β cell proliferation after short term High fat diet (HFD) treatment and
upregulation of phospho-S6K (downstream target of mTORC1) and Cyclin D2 in
in vitro INS-1 cell line treated with Palmitic Acid (PA). However, we do not know
the exact mechanism of activation. Although, there are no current studies that
can support our results in β cells, a recent study by Menon et al done in cancer
cell lines showed that mTOR can respond to the lipids via Phosphatidic acid de
novo synthesis [124]. This study utilized several cancer cell lines to demonstrate
the impact of exogenous saturated and unsaturated lipids on mTORC1 and
mTORC2. Their results indicated robust activation of both mTORC1 and
mTORC2 pathways, determined as the increase in phosphorylation levels of S6K
and AKT respectively, after 30 mins of treatment with oleic acid (unsaturated fatty
acid). Phosphatidic Acid has been previously reported to activate mTOR activity
[125, 126]. This study suggested that oleic acid gets incorporated into
Phosphatidic acid by LPAAT-β (Lysophosphatidic Acid Acyltransferase-β)
enzyme and activates mTOR. Though the authors saw modest activation of
mTORC1 by Palmitic Acid, they did not pursue it further. In summary, this study
suggested that exogenous lipids can get incorporated into Phosphatidic Acid that
can activate mTOR signaling pathways.

VI-1-2 Proposed Experiments
Since our study already showed that PA can activate mTORC1 pathway,
we next want to examine how fatty acids interact with mTOR. In order to
78
determine if mTOR interacts directly with fatty acids such as PA, we would study
if PA treatment displaces any of the proteins bound to mTORC1 complex and if
that displacement causes the activation of the pathway. mTORC1 complex is
made up of several proteins including mTOR itself, regulatory-associated protein
of mTOR (Raptor), mammalian lethal with SEC13 protein 8 (MLST8) and recently
identified PRAS40 and DEPTOR (Figure 25). Immunoprecipitation (IP)
experiments with anti-raptor antibody, with and without PA treatment, can help us
to determine evaluate if the association between the mTOR complex and Raptor
changes with PA and if that is downstream responsible for phosphorylation of
S6K. Additionally, since DEPTOR is an inhibitor of mTOR complex and is known
to bind to mTOR in the absence of mitogenic signal, activation of mTORC1
pathway would also indicate displacement of DEPTOR from mTOR complex. IP
experiments could also help us evaluate levels of DEPTOR associated with
mTORC1 complex, with and without PA treatment. Since our PA treatment
caused activation of downstream S6Kinase, we expect to see less association of
DEPTOR with mTORC1 complex after PA treatment.

VI-2 To determine if AKT activation can rescue the anti-proliferative effect
of long term activation of mTOR signaling on β cells.
Our in vitro results also indicated downregulation of AKT, determined by
studying the levels of phosphorylation level of AKT, in PA treated INS-1 cells
accompanied by upregulation of mTOR signaling. We also observed higher
levels of cleaved caspase 3, indicative of activation of apoptotic pathways in PA
79
treated cells. These results indicate the possible presence of a negative
feedback signaling taking place through mTOR pathway that is affecting AKT
signaling and ultimately influencing β cell proliferation (Figure 25). Therefore, we
would next want to study if AKT activation can rescue the anti-proliferative effect
of long term activation of mTOR signaling on β cells.

VI-2-1 Background & Rationale-
Insulin signaling has been seen to modulate pancreatic islet mass. Insulin
and insulin-like growth factor-1 (IGF-1) activate the signaling pathway by
stimulating Insulin receptor (tyrosine kinase receptors) and downstream insulin
receptor substrate (IRS). Out of the 6 members of the IRS family, IRS-2 is the
most functionally prominent in β cells and has been reported to control cell
growth and survival. While overexpression of IRS-2 has been shown to promote
mitogenesis [44, 127], deletion models lead to apoptosis and decreased β cell
survival leading to Diabetes [128-130]. β cell mass maintenance is critical in Type
2 Diabetic patients who experience loss in β cells along with increased
requirement of insulin, that together leads to hyperglycemia.
In insulin-responsive tissues such as skeletal muscle, it has been
observed that if mTOR is chronically activated (for example-due to increased
fatty acids), it can cause Serine/Threonine phosphorylation of IRS-1 and IRS-2,
leading to their ubiquitination and eventually degradation [109, 111].
Consequently, reducing insulin signaling in these tissues and leading to insulin
resistance. Similar results have been shown in β cells where prolonged activation
80
of mTOR signaling lead to Serine/Threonine phosphorylation of IRS-2 and cause
detrimental effects on β cell survival. A study by Briaud et al [131] in INS-1 cells
demonstrated that chronic activation of mTOR pathway by glucose and/or IGF
treatment for 24 hours led to higher phosphorylation levels of IRS-2, which
mediated its proteasomal degradation. Lower levels of IRS-2 were associated
with decreased phosphorylation of AKT and higher caspase 9 levels indicating
increased β cell apoptosis. This was reversed by using Kinase Dead (KD) mutant
of mTOR and mTOR inhibitor (Rapamycin), indicating the involvement of mTOR
pathway. Additionally, an in vivo study by Shigeyama et al showed that
prolonged activation of mTOR signaling in mice by deletion of TSC2 (mTOR
inhibitor) specifically in pancreatic β cells, led to hyperglycemia and decreased
islet mass & β cell number at 40 weeks of age [27]. These mice islets showed
decreased levels of active AKT and enhanced cleaved caspase-3 expression
along with reduced levels of IRS 1&2, indicating reduce insulin signaling and
enhanced cell apoptosis. To confirm the involvement of mTOR, βTSC2
-/-
mice
were treated with Rapamycin. Rapamycin treated mice did not develop
hyperglycemia, had no decrease in β cell mass and their islets showed enhanced
insulin signaling as indicated by higher levels of p-AKT and IRS-1&2, strongly
suggesting that mTORC1 activation was responsible. Furthermore, this study
looked at levels of phopho-S6K and phospho-4EBP1 proteins, indicative of
mTOR pathway activation, in mice treated with High Fat diet (HFD) for 3 weeks
and in 8-week old diabetic db/db mice. In both groups of mice, they observed
higher levels of phopho-S6K and phospho-4EBP1, supporting elevated mTOR
81
activity in these models of obesity and diabetes. In summary, this study reported
that chronic activation of mTOR signaling in mice causes decrease in insulin
signaling and activation of cellular apoptosis, together causing reduced β cell
mass as seen in Type 2 Diabetes.
Another study considering the role of mTORC1 and mTORC2 in Type 2
Diabetes pathology, revealed that mTORC1 pathway is highly activated in β cells
of diabetic individuals, while mTORC2 levels were greatly reduced [132].
Treatment of islets and INS-1 cell line with 22.2mM glucose for 72 hours (to
recreate hyperglycemic conditions) provided similar results, as indicated by
increased phosphorylation of S6K and 4EBP1 and reduced expression of
phospho-AKT. Selective inhibition of S6K in islets from Diabetic individuals
restored insulin secretion in these islets and enhanced AKT signaling indicative
of restored mTORC2 function. These results suggested that increased mTORC1
activity and decreased mTORC2 function are potential hallmarks on Type 2
Diabetes pathology.
82

Figure 25- Schematic diagram showing mTOR’s negative feedback
signaling to IRS and AKT

Together all the above-mentioned studies support, hyper-activation of
mTORC1 signaling under obesity and diabetic conditions, which sends a
negative feedback to the insulin signaling and causes decline in phospho-AKT
levels, which in turn have deleterious effects on β cell survival.

VI-2-2 Proposed Experiments
In order to elucidate whether replacement of active AKT levels in β cells
treated with PA will reverse/rescue the harmful effect of mTORC1 activation, we
would use 2 approaches-1.) PTEN is the negative regulator of the PI3K/AKT
signaling cascade, therefore deleting PTEN would cause higher levels of AKT in
the cell. Hence, we would knockdown PTEN in the β cells and look at the effect
of AKT activation on caspase 3 expression along with cell cycle inhibitory
83
proteins such as p16 and p27 in presence of fatty acids. If the restoration of AKT
levels can rescue the cells, we would see decreased levels of caspase 3, p16
and p27 in β cells with PTEN knock down as compared to untreated cells; 2.)
Secondly, we would transfect β cells with constitutive active (CA) form of AKT
and study the activation of apoptotic pathways along with the levels p16 and p27
using western blot in cells expressing CA-AKT in presence of fatty acids. Again, if
the higher levels of AKT is able to rescue the effect of prolonged mTORC1
activation, we would see lower levels of caspase 3, p16 and p27 expression in
CA-AKT expressing cells as compared to controls.

84
Chapter VII: Materials and Methods

Animals
Mice with two different background were used for this study- C57Bl/J6 and
Balb/c [133], since different backgrounds have been seen to have different
response to high fat diet. All animals were housed in a temperature-, humidity-,
light-controlled room (12-h light/dark cycle), allowing free access to food and
water. All experiments were conducted according to the Institutional Animal Care
and Use Committee of the University of Southern California research guidelines.

Diet Feeding
For the high fat diet (HFD) experiment, one group of mice were fed High
Fat Diet with 60 kcal% fat (TD06414, Harlan laboratories) whereas the control
group was fed Normal chow with13 kcal% of fat in their diet (PicoLab 5053)
(Table). Diet was started at 3 months of age and continued for 2 weeks, 2
months, 9 months and 12-14 months. Body weight, food intake and fed plasma
glucose levels were measured weekly.

Glucose Homeostasis
Blood glucose was determined by glucometer, using few microliters blood
from the tail vein. Intraperitoneal glucose (GTT) and Insulin (ITT) tolerance tests
were performed at the end of the study for 14 days and 2 months’ study and
every alternate month for 9 months’ and 12-14 months’ treatment. GTT was
performed in 16-17 hours fasted mice by injecting 2g/Kg D-glucose
85
intraperitoneally while ITT was done in 4-6 hours fasted mice by injecting
0.5U/Kg Novolin R (human insulin).

Cell Culture
INS-1 cells were kindly supplied by Prasanna Dadi at Vanderbilt, TN and
cultured at 37 °C in a humidified atmosphere containing 5% CO2 in RPMI 1640
medium containing 11 mM glucose and supplemented with 10% heat-inactivated
FBS, 50 μM 2-mercaptoethanol, 100 units/ml penicillin and 100 μg/ml
streptomycin. Cells were starved overnight 48 hours after seeding and then
treated with fatty acids in RPMI media containing 6mM glucose without FBS.

Fatty Acid Preparation
Palmitic Acid (sigma#P0500-10G) 200mM stock was prepared by
dissolving 51.2mg PA in 1ml 100% ethanol. 0.04ml of this stock was dissolved in
1.96ml 10% fatty acid free BSA (in DMEM media) for 4mM PA stock by shaking
overnight at room temperature. Solution was filtered and was diluted 1:10 times
prior to cell treatment. 1% BSA was used as a control.

Biochemical Assays
Insulin Assay- Mouse Ultrasensitive Insulin ELISA kit was used from
ALPCO (Cat#80-INSMSU-E01) and plasma Insulin was quantified as per kit
instructions.
NEFA (non-esterified fatty acid)/ Triglyceride Assay- Wako NEFA and
86
Triglyceride quantification kits were used to quantify plasma fatty acid and
triglyceride levels respectively.

Mouse islet isolation
Pancreases were perfused with collagenase P solution (0.8 mg/ml; 5ml
per mouse) and digested at 37 ºC for 17 min. Islets were then purified by using
Ficoll gradients with densities of 1.108, 1.096, 1.069 and 1.037 (Cellgro) as
previously reported [134].

Immunohistochemistry
Zn-formalin fixed and paraffin embedded sections were stained as
previously reported [134]. Antibodies used are: Cyclin D1 (Santa Cruz, sc-8396),
Cyclin D2 (Santa Cruz, sc-593), Ki-67 monoclonal Ab (CST#12202), p27 (Santa
Cruz, sc-1641), BrDU.

Western blot
Cell lysate preparation and immunoblot analysis were performed as
described [135]. Briefly, cells or islets were lysed in cell lysis buffer. Supernatants
of the lysates were subjected to SDS-PAGE (10-12% polyacrylamide gel) and
then transferred to PVDF membranes. Antibodies used: Phopho-4E-BP1 (Ser65)
(CST #9451), Phospho-p70 S6 Kinase (Thr389) (CST#9205), Phospho-p44/42
MAPK (Erk1/2) (Thr202/Tyr204) (CST#4370), Cleaved Caspase-3 (Asp175)
(CST#9661), Cyclin A (Santa Cruz, sc-751), Cyclin D1 (Santa Cruz, sc-8396),
87
Cyclin D2 (Santa Cruz, sc-593), Ki-67 monoclonal Ab (CST#12202), p27 (Santa
Cruz, sc-1641). ECL Secondary Mouse and Rabbit HRP antibodies used were
from GE Healthcare.

MTT Assay
Cells were seeded at density 1.5-2x10
4
cell/well in 96-well plates in RPMI
media. After 48 hours of seeding, the cells were treated with 1% BSA (as control)
or different concentrations of Palmitic Acid (0.2mM, 0.4mM, 0.6mM, 0.8mM and
1mM) (6 wells per treatment) and incubated for 24 and 48 hours. 10ul MTT
reagent was then added after respective incubation and kept at 37C for 4 hours,
followed by addition of 100ul DMSO. Plate was kept on shaker for 5-10 minutes
to dissolve the crystals. Optical Density Reading was then taken at 570 nm
wavelength.

Growth Curve
Cells were seeded at density 5-6x10
4
cell/well in six-well plates in RPMI
media. After 48 hours, plate 1 was washed and cells were trypsinized & counted.
This was considered day 0. Rest of the plates were treated with BSA or PA (in
triplicates) and incubated for 1, 3, 5, 8 days. Cells were washed, trypsinized and
counted after their respective incubation. The results were plotted as a graph on
excel.

88
Cell Cycle FACS
INS-1 cells (3x10
5
cells/well) were seeded in six-well plates in RPMI media
with 6mM glucose with FBS. Starved cells for 48-72 hours in media without FBS
after 24 hours of seeding. They were then treated with 1% BSA or 0.4mM
Palmitic Acid for 48 hours. Cells were harvested by washing, trypsinizing and
centrifuging them. The resulting pellet was re-suspended in 0.1ml PBS. 1ml
ethanol was then added (kept at -20C) and kept for 20 mins at -20C. Cells were
centrifuged at 1000rpm for 5 mins and supernatant was discarded. Then the
pellet was re-suspended in 500ul of RNase solution (200ug/ml in PBS) and
incubated at room temperature for 30 mins. Propidium Iodide was later added to
the mix (50ug/ml) and incubated for 30 mins away from light. FACS analysis was
performed using FACS calibur machine and software.

PI/ANNEXIN V FACS
INS-1 cells were seeded at density (2.5x10
5
cells/well) in six-well plates in
RPMI media. After 48 hours of seeding, cells were treated with 1% BSA or
0.4mM Palmitic Acid for 48 hours. Media containing detached and floating cells
was collected and the rest of the cells were trypsinized, and washed once with
PBS. Cells were then washed and suspended in 1X Annexin Binding Buffer
(ABB). Diluted Annexin (400ng per 1x10
6
cells) was then added to cells and
incubated for 8 minutes at room temperature away from light. Propidium Iodide
was then added to each sample (2.5ug/ml per sample) and incubated for
approximately 2 minutes. FACS analysis was then performed.
89

Rapamycin treatment in mice
Mice were fed HFD for the required duration and rapamycin was injected
intraperitoneally on the last 8 days of treatment (everyday 0.3mg/Kg per mouse).
100mM Rapamycin stock in DMSO (LC labs # R-5000) was diluted 100 times by
mixing 890ul PBS, 100ul Tween 80 with 10ul rapamycin stock. Calculations were
then done and mice were injected as per their body weight.

BrDU treatment
BrDU (1mg/ml concentration) was given to mice in water for 5 days before
ending the study. Mice were euthanized after the 5 days and organs were
collected as per the experiment.

RNA sequencing and Data Analysis
Six mice islet preparations (3 in Normal chow group and 3 in HFD group)
were sequenced and data was analyzed using Partek Flow and IPA (Ingenuity
Pathway Analysis). In brief, RNeasy Mini Kit (Qiagen, Cat# 74104) was used to
isolate total RNA from mice islets. RNA quality was tested using Agilent
Bioanalyzer and the RNA integrity number (RIN) values for all the samples were
>7.5. We utilized USC NGS core services to convert mRNA to cDNA libraries
which was further sequenced using Illumina NextSeq500- 25 million reads per
sample.

90
Statistical Analysis
The data are presented as means ± the standard error of the mean
(SEM). Differences between individual groups were analyzed by Student’s t test,
with two-tailed p values less than 0.05 considered statistically significant.

Table 1. Diet composition table

Kcal% Normal Chow (NC) High Fat Diet (HFD)
CHO 62 21
Fat 13 60
Protein 24.5 80

91
Bibliography

[1] T.A. Buchanan, Pancreatic beta-cell loss and preservation in type 2 diabetes, Clin
Ther, 25 Suppl B (2003) B32-46.
[2] G. Gu, J. Dubauskaite, D.A. Melton, Direct evidence for the pancreatic lineage:
NGN3+ cells are islet progenitors and are distinct from duct progenitors,
Development, 129 (2002) 2447-2457.
[3] Y. Kawaguchi, B. Cooper, M. Gannon, M. Ray, R.J. MacDonald, C.V. Wright, The role
of the transcriptional regulator Ptf1a in converting intestinal to pancreatic
progenitors, Nat Genet, 32 (2002) 128-134.
[4] R.C. McEvoy, K.L. Madson, Pancreatic insulikn-, glucagon-, and somatostatin-
positive islet cell populatins during the perinatal development of the rat. I.
Morphometric quantitation, Biol Neonate, 38 (1980) 248-254.
[5] L. Bouwens, R.N. Wang, E. De Blay, D.G. Pipeleers, G. Kloppel, Cytokeratins as
markers of ductal cell differentiation and islet neogenesis in the neonatal rat
pancreas, Diabetes, 43 (1994) 1279-1283.
[6] L. Bouwens, I. Rooman, Regulation of pancreatic beta-cell mass, Physiol Rev, 85
(2005) 1255-1270.
[7] D.T. Finegood, L. Scaglia, S. Bonner-Weir, Dynamics of beta-cell mass in the
growing rat pancreas. Estimation with a simple mathematical model, Diabetes, 44
(1995) 249-256.
[8] M. Teta, S.Y. Long, L.M. Wartschow, M.M. Rankin, J.A. Kushner, Very slow
turnover of beta-cells in aged adult mice, Diabetes, 54 (2005) 2557-2567.
[9] A.M. Ackermann, M. Gannon, Molecular regulation of pancreatic beta-cell mass
development, maintenance, and expansion, J Mol Endocrinol, 38 (2007) 193-206.
[10] Y. Dor, J. Brown, O.I. Martinez, D.A. Melton, Adult pancreatic beta-cells are
formed by self-duplication rather than stem-cell differentiation, Nature, 429 (2004)
41-46.
[11] R.N. Kulkarni, J.C. Bruning, J.N. Winnay, C. Postic, M.A. Magnuson, C.R. Kahn,
Tissue-specific knockout of the insulin receptor in pancreatic beta cells creates an
insulin secretory defect similar to that in type 2 diabetes, Cell, 96 (1999) 329-339.
[12] R.N. Kulkarni, M. Holzenberger, D.Q. Shih, U. Ozcan, M. Stoffel, M.A. Magnuson,
C.R. Kahn, beta-cell-specific deletion of the Igf1 receptor leads to hyperinsulinemia
and glucose intolerance but does not alter beta-cell mass, Nat Genet, 31 (2002) 111-
115.
[13] K. Otani, R.N. Kulkarni, A.C. Baldwin, J. Krutzfeldt, K. Ueki, M. Stoffel, C.R. Kahn,
K.S. Polonsky, Reduced beta-cell mass and altered glucose sensing impair insulin-
secretory function in betaIRKO mice, Am J Physiol Endocrinol Metab, 286 (2004)
E41-49.
[14] C. Dai, C.G. Huh, S.S. Thorgeirsson, Y. Liu, Beta-cell-specific ablation of the
hepatocyte growth factor receptor results in reduced islet size, impaired insulin
secretion, and glucose intolerance, Am J Pathol, 167 (2005) 429-436.
[15] A. Garcia-Ocana, K.K. Takane, M.A. Syed, W.M. Philbrick, R.C. Vasavada, A.F.
Stewart, Hepatocyte growth factor overexpression in the islet of transgenic mice
92
increases beta cell proliferation, enhances islet mass, and induces mild
hypoglycemia, J Biol Chem, 275 (2000) 1226-1232.
[16] W. Liu, C. Chin-Chance, E.J. Lee, W.L. Lowe, Jr., Activation of
phosphatidylinositol 3-kinase contributes to insulin-like growth factor I-mediated
inhibition of pancreatic beta-cell death, Endocrinology, 143 (2002) 3802-3812.
[17] R.L. Tuttle, N.S. Gill, W. Pugh, J.P. Lee, B. Koeberlein, E.E. Furth, K.S. Polonsky, A.
Naji, M.J. Birnbaum, Regulation of pancreatic beta-cell growth and survival by the
serine/threonine protein kinase Akt1/PKBalpha, Nat Med, 7 (2001) 1133-1137.
[18] E. Bernal-Mizrachi, W. Wen, S. Stahlhut, C.M. Welling, M.A. Permutt, Islet beta
cell expression of constitutively active Akt1/PKB alpha induces striking
hypertrophy, hyperplasia, and hyperinsulinemia, J Clin Invest, 108 (2001) 1631-
1638.
[19] S. Fatrai, L. Elghazi, N. Balcazar, C. Cras-Meneur, I. Krits, H. Kiyokawa, E. Bernal-
Mizrachi, Akt induces beta-cell proliferation by regulating cyclin D1, cyclin D2, and
p21 levels and cyclin-dependent kinase-4 activity, Diabetes, 55 (2006) 318-325.
[20] K. Inoki, Y. Li, T. Zhu, J. Wu, K.L. Guan, TSC2 is phosphorylated and inhibited by
Akt and suppresses mTOR signalling, Nat Cell Biol, 4 (2002) 648-657.
[21] D.D. Sarbassov, S.M. Ali, D.H. Kim, D.A. Guertin, R.R. Latek, H. Erdjument-
Bromage, P. Tempst, D.M. Sabatini, Rictor, a novel binding partner of mTOR, defines
a rapamycin-insensitive and raptor-independent pathway that regulates the
cytoskeleton, Curr Biol, 14 (2004) 1296-1302.
[22] D.D. Sarbassov, D.A. Guertin, S.M. Ali, D.M. Sabatini, Phosphorylation and
regulation of Akt/PKB by the rictor-mTOR complex, Science, 307 (2005) 1098-1101.
[23] E. Jacinto, R. Loewith, A. Schmidt, S. Lin, M.A. Ruegg, A. Hall, M.N. Hall,
Mammalian TOR complex 2 controls the actin cytoskeleton and is rapamycin
insensitive, Nat Cell Biol, 6 (2004) 1122-1128.
[24] H. Shima, M. Pende, Y. Chen, S. Fumagalli, G. Thomas, S.C. Kozma, Disruption of
the p70(s6k)/p85(s6k) gene reveals a small mouse phenotype and a new functional
S6 kinase, EMBO J, 17 (1998) 6649-6659.
[25] M. Pende, S.C. Kozma, M. Jaquet, V. Oorschot, R. Burcelin, Y. Le Marchand-
Brustel, J. Klumperman, B. Thorens, G. Thomas, Hypoinsulinaemia, glucose
intolerance and diminished beta-cell size in S6K1-deficient mice, Nature, 408 (2000)
994-997.
[26] L. Rachdi, N. Balcazar, F. Osorio-Duque, L. Elghazi, A. Weiss, A. Gould, K.J. Chang-
Chen, M.J. Gambello, E. Bernal-Mizrachi, Disruption of Tsc2 in pancreatic beta cells
induces beta cell mass expansion and improved glucose tolerance in a TORC1-
dependent manner, Proc Natl Acad Sci U S A, 105 (2008) 9250-9255.
[27] Y. Shigeyama, T. Kobayashi, Y. Kido, N. Hashimoto, S. Asahara, T. Matsuda, A.
Takeda, T. Inoue, Y. Shibutani, M. Koyanagi, T. Uchida, M. Inoue, O. Hino, M. Kasuga,
T. Noda, Biphasic response of pancreatic beta-cell mass to ablation of tuberous
sclerosis complex 2 in mice, Mol Cell Biol, 28 (2008) 2971-2979.
[28] E. Zahr, R.D. Molano, A. Pileggi, H. Ichii, S.S. Jose, N. Bocca, W. An, J. Gonzalez-
Quintana, C. Fraker, C. Ricordi, L. Inverardi, Rapamycin impairs in vivo proliferation
of islet beta-cells, Transplantation, 84 (2007) 1576-1583.
[29] N. Balcazar, A. Sathyamurthy, L. Elghazi, A. Gould, A. Weiss, I. Shiojima, K.
Walsh, E. Bernal-Mizrachi, mTORC1 activation regulates beta-cell mass and
93
proliferation by modulation of cyclin D2 synthesis and stability, J Biol Chem, 284
(2009) 7832-7842.
[30] S. Georgia, A. Bhushan, Beta cell replication is the primary mechanism for
maintaining postnatal beta cell mass, J Clin Invest, 114 (2004) 963-968.
[31] J.A. Kushner, M.A. Ciemerych, E. Sicinska, L.M. Wartschow, M. Teta, S.Y. Long, P.
Sicinski, M.F. White, Cyclins D2 and D1 are essential for postnatal pancreatic beta-
cell growth, Mol Cell Biol, 25 (2005) 3752-3762.
[32] I. Cozar-Castellano, K.K. Takane, R. Bottino, A.N. Balamurugan, A.F. Stewart,
Induction of beta-cell proliferation and retinoblastoma protein phosphorylation in
rat and human islets using adenovirus-mediated transfer of cyclin-dependent
kinase-4 and cyclin D1, Diabetes, 53 (2004) 149-159.
[33] J. Krishnamurthy, M.R. Ramsey, K.L. Ligon, C. Torrice, A. Koh, S. Bonner-Weir,
N.E. Sharpless, p16INK4a induces an age-dependent decline in islet regenerative
potential, Nature, 443 (2006) 453-457.
[34] S. Georgia, A. Bhushan, p27 Regulates the transition of beta-cells from
quiescence to proliferation, Diabetes, 55 (2006) 2950-2956.
[35] T. Uchida, T. Nakamura, N. Hashimoto, T. Matsuda, K. Kotani, H. Sakaue, Y. Kido,
Y. Hayashi, K.I. Nakayama, M.F. White, M. Kasuga, Deletion of Cdkn1b ameliorates
hyperglycemia by maintaining compensatory hyperinsulinemia in diabetic mice, Nat
Med, 11 (2005) 175-182.
[36] L. Rachdi, N. Balcazar, L. Elghazi, D.J. Barker, I. Krits, H. Kiyokawa, E. Bernal-
Mizrachi, Differential effects of p27 in regulation of beta-cell mass during
development, neonatal period, and adult life, Diabetes, 55 (2006) 3520-3528.
[37] C. Bernard, M.F. Berthault, C. Saulnier, A. Ktorza, Neogenesis vs. apoptosis As
main components of pancreatic beta cell ass changes in glucose-infused normal and
mildly diabetic adult rats, FASEB J, 13 (1999) 1195-1205.
[38] B.G. Topp, M.D. McArthur, D.T. Finegood, Metabolic adaptations to chronic
glucose infusion in rats, Diabetologia, 47 (2004) 1602-1610.
[39] K. Maedler, D.M. Schumann, F. Schulthess, J. Oberholzer, D. Bosco, T. Berney,
M.Y. Donath, Aging correlates with decreased beta-cell proliferative capacity and
enhanced sensitivity to apoptosis: a potential role for Fas and pancreatic duodenal
homeobox-1, Diabetes, 55 (2006) 2455-2462.
[40] F.M. Matschinsky, Assessing the potential of glucokinase activators in diabetes
therapy, Nat Rev Drug Discov, 8 (2009) 399-416.
[41] Y. Terauchi, H. Sakura, K. Yasuda, K. Iwamoto, N. Takahashi, K. Ito, H. Kasai, H.
Suzuki, O. Ueda, N. Kamada, et al., Pancreatic beta-cell-specific targeted disruption of
glucokinase gene. Diabetes mellitus due to defective insulin secretion to glucose, J
Biol Chem, 270 (1995) 30253-30256.
[42] Y. Terauchi, I. Takamoto, N. Kubota, J. Matsui, R. Suzuki, K. Komeda, A. Hara, Y.
Toyoda, I. Miwa, S. Aizawa, S. Tsutsumi, Y. Tsubamoto, S. Hashimoto, K. Eto, A.
Nakamura, M. Noda, K. Tobe, H. Aburatani, R. Nagai, T. Kadowaki, Glucokinase and
IRS-2 are required for compensatory beta cell hyperplasia in response to high-fat
diet-induced insulin resistance, J Clin Invest, 117 (2007) 246-257.
[43] N. Kubota, K. Tobe, Y. Terauchi, K. Eto, T. Yamauchi, R. Suzuki, Y. Tsubamoto, K.
Komeda, R. Nakano, H. Miki, S. Satoh, H. Sekihara, S. Sciacchitano, M. Lesniak, S.
Aizawa, R. Nagai, S. Kimura, Y. Akanuma, S.I. Taylor, T. Kadowaki, Disruption of
94
insulin receptor substrate 2 causes type 2 diabetes because of liver insulin
resistance and lack of compensatory beta-cell hyperplasia, Diabetes, 49 (2000)
1880-1889.
[44] A.M. Hennige, D.J. Burks, U. Ozcan, R.N. Kulkarni, J. Ye, S. Park, M. Schubert, T.L.
Fisher, M.A. Dow, R. Leshan, M. Zakaria, M. Mossa-Basha, M.F. White, Upregulation of
insulin receptor substrate-2 in pancreatic beta cells prevents diabetes, J Clin Invest,
112 (2003) 1521-1532.
[45] V. Poitout, R.P. Robertson, Glucolipotoxicity: fuel excess and beta-cell
dysfunction, Endocr Rev, 29 (2008) 351-366.
[46] J.W. Kim, K.H. Yoon, Glucolipotoxicity in Pancreatic beta-Cells, Diabetes Metab J,
35 (2011) 444-450.
[47] V. Poitout, J. Amyot, M. Semache, B. Zarrouki, D. Hagman, G. Fontes,
Glucolipotoxicity of the pancreatic beta cell, Biochim Biophys Acta, 1801 (2010)
289-298.
[48] K. Maedler, J. Oberholzer, P. Bucher, G.A. Spinas, M.Y. Donath, Monounsaturated
fatty acids prevent the deleterious effects of palmitate and high glucose on human
pancreatic beta-cell turnover and function, Diabetes, 52 (2003) 726-733.
[49] T.C. Brelje, N.V. Bhagroo, L.E. Stout, R.L. Sorenson, Beneficial effects of lipids
and prolactin on insulin secretion and beta-cell proliferation: a role for lipids in the
adaptation of islets to pregnancy, J Endocrinol, 197 (2008) 265-276.
[50] Z.Y. Chen, J. Yang, S.C. Cunnane, Gestational hyperlipidemia in the rat is
characterized by accumulation of n - 6 and n - 3 fatty acids, especially
docosahexaenoic acid, Biochim Biophys Acta, 1127 (1992) 263-269.
[51] R.H. Knopp, C.D. Saudek, R.A. Arky, J.B. O'Sullivan, 2 phases of adipose tissue
metabolism in pregnancy: maternal adaptations for fetal growth, Endocrinology, 92
(1973) 984-988.
[52] J.A. Gonzalez-Pertusa, J. Dube, S.R. Valle, T.C. Rosa, K.K. Takane, J.M. Mellado-Gil,
G. Perdomo, R.C. Vasavada, A. Garcia-Ocana, Novel proapoptotic effect of hepatocyte
growth factor: synergy with palmitate to cause pancreatic {beta}-cell apoptosis,
Endocrinology, 151 (2010) 1487-1498.
[53] J.L. Milburn, Jr., H. Hirose, Y.H. Lee, Y. Nagasawa, A. Ogawa, M. Ohneda, H.
BeltrandelRio, C.B. Newgard, J.H. Johnson, R.H. Unger, Pancreatic beta-cells in
obesity. Evidence for induction of functional, morphologic, and metabolic
abnormalities by increased long chain fatty acids, J Biol Chem, 270 (1995) 1295-
1299.
[54] Y.P. Zhou, V.E. Grill, Long-term exposure of rat pancreatic islets to fatty acids
inhibits glucose-induced insulin secretion and biosynthesis through a glucose fatty
acid cycle, J Clin Invest, 93 (1994) 870-876.
[55] A. Pick, J. Clark, C. Kubstrup, M. Levisetti, W. Pugh, S. Bonner-Weir, K.S.
Polonsky, Role of apoptosis in failure of beta-cell mass compensation for insulin
resistance and beta-cell defects in the male Zucker diabetic fatty rat, Diabetes, 47
(1998) 358-364.
[56] J. Pascoe, D. Hollern, R. Stamateris, M. Abbasi, L.C. Romano, B. Zou, C.P.
O'Donnell, A. Garcia-Ocana, L.C. Alonso, Free fatty acids block glucose-induced beta-
cell proliferation in mice by inducing cell cycle inhibitors p16 and p18, Diabetes, 61
(2012) 632-641.
95
[57] M. Prentki, B.E. Corkey, Are the beta-cell signaling molecules malonyl-CoA and
cystolic long-chain acyl-CoA implicated in multiple tissue defects of obesity and
NIDDM?, Diabetes, 45 (1996) 273-283.
[58] G. Solinas, W. Naugler, F. Galimi, M.S. Lee, M. Karin, Saturated fatty acids inhibit
induction of insulin gene transcription by JNK-mediated phosphorylation of insulin-
receptor substrates, Proc Natl Acad Sci U S A, 103 (2006) 16454-16459.
[59] S. Bonner-Weir, L.A. Baxter, G.T. Schuppin, F.E. Smith, A second pathway for
regeneration of adult exocrine and endocrine pancreas. A possible recapitulation of
embryonic development, Diabetes, 42 (1993) 1715-1720.
[60] G. Zajicek, N. Arber, D. Schwartz-Arad, I. Ariel, Streaming pancreas: islet cell
kinetics, Diabetes Res, 13 (1990) 121-125.
[61] H. Zulewski, E.J. Abraham, M.J. Gerlach, P.B. Daniel, W. Moritz, B. Muller, M.
Vallejo, M.K. Thomas, J.F. Habener, Multipotential nestin-positive stem cells isolated
from adult pancreatic islets differentiate ex vivo into pancreatic endocrine, exocrine,
and hepatic phenotypes, Diabetes, 50 (2001) 521-533.
[62] M. Lipsett, D.T. Finegood, beta-cell neogenesis during prolonged hyperglycemia
in rats, Diabetes, 51 (2002) 1834-1841.
[63] S. Bonner-Weir, Life and death of the pancreatic beta cells, Trends Endocrinol
Metab, 11 (2000) 375-378.
[64] S.I. Tschen, S. Dhawan, T. Gurlo, A. Bhushan, Age-dependent decline in beta-cell
proliferation restricts the capacity of beta-cell regeneration in mice, Diabetes, 58
(2009) 1312-1320.
[65] R.S. Surwit, C.M. Kuhn, C. Cochrane, J.A. McCubbin, M.N. Feinglos, Diet-induced
type II diabetes in C57BL/6J mice, Diabetes, 37 (1988) 1163-1167.
[66] M.S. Winzell, B. Ahren, The high-fat diet-fed mouse: a model for studying
mechanisms and treatment of impaired glucose tolerance and type 2 diabetes,
Diabetes, 53 Suppl 3 (2004) S215-219.
[67] R.E. Stamateris, R.B. Sharma, D.A. Hollern, L.C. Alonso, Adaptive beta-cell
proliferation increases early in high-fat feeding in mice, concurrent with metabolic
changes, with induction of islet cyclin D2 expression, Am J Physiol Endocrinol
Metab, 305 (2013) E149-159.
[68] H. Sone, Y. Kagawa, Pancreatic beta cell senescence contributes to the
pathogenesis of type 2 diabetes in high-fat diet-induced diabetic mice, Diabetologia,
48 (2005) 58-67.
[69] R.L. Hull, K. Kodama, K.M. Utzschneider, D.B. Carr, R.L. Prigeon, S.E. Kahn,
Dietary-fat-induced obesity in mice results in beta cell hyperplasia but not increased
insulin release: evidence for specificity of impaired beta cell adaptation,
Diabetologia, 48 (2005) 1350-1358.
[70] S. Georgia, C. Hinault, D. Kawamori, J. Hu, J. Meyer, M. Kanji, A. Bhushan, R.N.
Kulkarni, Cyclin D2 is essential for the compensatory beta-cell hyperplastic
response to insulin resistance in rodents, Diabetes, 59 (2010) 987-996.
[71] S.J. Salpeter, A. Klochendler, N. Weinberg-Corem, S. Porat, Z. Granot, A.M.
Shapiro, M.A. Magnuson, A. Eden, J. Grimsby, B. Glaser, Y. Dor, Glucose regulates
cyclin D2 expression in quiescent and replicating pancreatic beta-cells through
glycolysis and calcium channels, Endocrinology, 152 (2011) 2589-2598.
96
[72] R.E. Stamateris, R.B. Sharma, Y. Kong, P. Ebrahimpour, D. Panday, P. Ranganath,
B. Zou, H. Levitt, N.A. Parambil, C.P. O'Donnell, A. Garcia-Ocana, L.C. Alonso, Glucose
Induces Mouse beta-Cell Proliferation via IRS2, MTOR, and Cyclin D2 but Not the
Insulin Receptor, Diabetes, 65 (2016) 981-995.
[73] A.E. Butler, J. Janson, S. Bonner-Weir, R. Ritzel, R.A. Rizza, P.C. Butler, Beta-cell
deficit and increased beta-cell apoptosis in humans with type 2 diabetes, Diabetes,
52 (2003) 102-110.
[74] C.J. Rhodes, IGF-I and GH post-receptor signaling mechanisms for pancreatic
beta-cell replication, J Mol Endocrinol, 24 (2000) 303-311.
[75] J.H. Nielsen, E.D. Galsgaard, A. Moldrup, B.N. Friedrichsen, N. Billestrup, J.A.
Hansen, Y.C. Lee, C. Carlsson, Regulation of beta-cell mass by hormones and growth
factors, Diabetes, 50 Suppl 1 (2001) S25-29.
[76] C.J. Nolan, M.S. Madiraju, V. Delghingaro-Augusto, M.L. Peyot, M. Prentki, Fatty
acid signaling in the beta-cell and insulin secretion, Diabetes, 55 Suppl 2 (2006) S16-
23.
[77] R.E. Mosser, M.F. Maulis, V.S. Moulle, J.C. Dunn, B.A. Carboneau, K. Arasi, K.
Pappan, V. Poitout, M. Gannon, High-fat diet-induced beta-cell proliferation occurs
prior to insulin resistance in C57Bl/6J male mice, Am J Physiol Endocrinol Metab,
308 (2015) E573-582.
[78] J. Ahren, B. Ahren, N. Wierup, Increased beta-cell volume in mice fed a high-fat
diet: a dynamic study over 12 months, Islets, 2 (2010) 353-356.
[79] M. Higa, M. Shimabukuro, Y. Shimajiri, N. Takasu, T. Shinjyo, T. Inaba, Protein
kinase B/Akt signalling is required for palmitate-induced beta-cell lipotoxicity,
Diabetes Obes Metab, 8 (2006) 228-233.
[80] L. Liu, R.P. Wang, X.H. Liu, L.X. Wang, X.Y. Liu, W.J. Chen, L.B. Liu, [The chronic
effect of palmitic acid on apoptosis of pancreatic islet beta-cells and the mechanism],
Zhongguo Ying Yong Sheng Li Xue Za Zhi, 25 (2009) 553-556.
[81] Q. Liu, R. Wang, H. Zhou, L. Zhang, Y. Cao, X. Wang, Y. Hao, SHIP2 on pI3K/Akt
pathway in palmitic acid stimulated islet beta cell, Int J Clin Exp Med, 8 (2015) 3210-
3218.
[82] I. Cozar-Castellano, M. Weinstock, M. Haught, S. Velazquez-Garcia, D. Sipula, A.F.
Stewart, Evaluation of beta-cell replication in mice transgenic for hepatocyte growth
factor and placental lactogen: comprehensive characterization of the G1/S
regulatory proteins reveals unique involvement of p21cip, Diabetes, 55 (2006) 70-
77.
[83] K.N. Frayn, C.M. Williams, P. Arner, Are increased plasma non-esterified fatty
acid concentrations a risk marker for coronary heart disease and other chronic
diseases?, Clin Sci (Lond), 90 (1996) 243-253.
[84] N. Turner, G.M. Kowalski, S.J. Leslie, S. Risis, C. Yang, R.S. Lee-Young, J.R. Babb,
P.J. Meikle, G.I. Lancaster, D.C. Henstridge, P.J. White, E.W. Kraegen, A. Marette, G.J.
Cooney, M.A. Febbraio, C.R. Bruce, Distinct patterns of tissue-specific lipid
accumulation during the induction of insulin resistance in mice by high-fat feeding,
Diabetologia, 56 (2013) 1638-1648.
[85] J.L. Jewell, K.L. Guan, Nutrient signaling to mTOR and cell growth, Trends
Biochem Sci, 38 (2013) 233-242.
97
[86] J. Kim, K.L. Guan, Amino acid signaling in TOR activation, Annu Rev Biochem, 80
(2011) 1001-1032.
[87] L. Wang, Y. Liu, S. Yan Lu, K.T. Nguyen, S.A. Schroer, A. Suzuki, T.W. Mak, H.
Gaisano, M. Woo, Deletion of Pten in pancreatic ss-cells protects against deficient ss-
cell mass and function in mouse models of type 2 diabetes, Diabetes, 59 (2010)
3117-3126.
[88] M. Cnop, B. Abdulkarim, G. Bottu, D.A. Cunha, M. Igoillo-Esteve, M. Masini, J.V.
Turatsinze, T. Griebel, O. Villate, I. Santin, M. Bugliani, L. Ladriere, L. Marselli, M.I.
McCarthy, P. Marchetti, M. Sammeth, D.L. Eizirik, RNA sequencing identifies
dysregulation of the human pancreatic islet transcriptome by the saturated fatty
acid palmitate, Diabetes, 63 (2014) 1978-1993.
[89] W. Zhang, H.T. Liu, MAPK signal pathways in the regulation of cell proliferation
in mammalian cells, Cell Res, 12 (2002) 9-18.
[90] J. Lakshmipathi, J.C. Alvarez-Perez, C. Rosselot, G.P. Casinelli, R.E. Stamateris, F.
Rausell-Palamos, C.P. O'Donnell, R.C. Vasavada, D.K. Scott, L.C. Alonso, A. Garcia-
Ocana, PKCzeta Is Essential for Pancreatic beta-Cell Replication During Insulin
Resistance by Regulating mTOR and Cyclin-D2, Diabetes, 65 (2016) 1283-1296.
[91] T. Hirai, K. Chida, Protein kinase Czeta (PKCzeta): activation mechanisms and
cellular functions, J Biochem, 133 (2003) 1-7.
[92] Y.F. Li, H.X. Sun, G.D. Wu, W.N. Du, J. Zuo, Y. Shen, B.Q. Qiang, Z.J. Yao, H. Wang,
W. Huang, Z. Chen, M.M. Xiong, Y. Meng, F.D. Fang, Protein kinase C/zeta (PRKCZ)
gene is associated with type 2 diabetes in Han population of North China and
analysis of its haplotypes, World J Gastroenterol, 9 (2003) 2078-2082.
[93] L. Qin, L. Zhou, X. Wu, J. Cheng, J. Wang, Y. Du, J. Hu, M. Shen, J. Zhou, Genetic
variants in protein kinase C zeta gene and type 2 diabetes risk: a case-control study
of a Chinese Han population, Diabetes Metab Res Rev, 24 (2008) 480-485.
[94] L.K. Olson, J.B. Redmon, H.C. Towle, R.P. Robertson, Chronic exposure of HIT
cells to high glucose concentrations paradoxically decreases insulin gene
transcription and alters binding of insulin gene regulatory protein, J Clin Invest, 92
(1993) 514-519.
[95] V. Poitout, L.K. Olson, R.P. Robertson, Chronic exposure of betaTC-6 cells to
supraphysiologic concentrations of glucose decreases binding of the RIPE3b1
insulin gene transcription activator, J Clin Invest, 97 (1996) 1041-1046.
[96] H. Kaneto, T.A. Matsuoka, Y. Nakatani, D. Kawamori, T. Miyatsuka, M. Matsuhisa,
Y. Yamasaki, Oxidative stress, ER stress, and the JNK pathway in type 2 diabetes, J
Mol Med (Berl), 83 (2005) 429-439.
[97] M. Shimabukuro, Y.T. Zhou, M. Levi, R.H. Unger, Fatty acid-induced beta cell
apoptosis: a link between obesity and diabetes, Proc Natl Acad Sci U S A, 95 (1998)
2498-2502.
[98] S. Piro, M. Anello, C. Di Pietro, M.N. Lizzio, G. Patane, A.M. Rabuazzo, R. Vigneri,
M. Purrello, F. Purrello, Chronic exposure to free fatty acids or high glucose induces
apoptosis in rat pancreatic islets: possible role of oxidative stress, Metabolism, 51
(2002) 1340-1347.
[99] R. Lupi, F. Dotta, L. Marselli, S. Del Guerra, M. Masini, C. Santangelo, G. Patane, U.
Boggi, S. Piro, M. Anello, E. Bergamini, F. Mosca, U. Di Mario, S. Del Prato, P.
Marchetti, Prolonged exposure to free fatty acids has cytostatic and pro-apoptotic
98
effects on human pancreatic islets: evidence that beta-cell death is caspase
mediated, partially dependent on ceramide pathway, and Bcl-2 regulated, Diabetes,
51 (2002) 1437-1442.
[100] J. Krishnamurthy, C. Torrice, M.R. Ramsey, G.I. Kovalev, K. Al-Regaiey, L. Su,
N.E. Sharpless, Ink4a/Arf expression is a biomarker of aging, J Clin Invest, 114
(2004) 1299-1307.
[101] F. Karpe, J.R. Dickmann, K.N. Frayn, Fatty acids, obesity, and insulin resistance:
time for a reevaluation, Diabetes, 60 (2011) 2441-2449.
[102] R.B. Sharma, L.C. Alonso, Lipotoxicity in the pancreatic beta cell: not just
survival and function, but proliferation as well?, Curr Diab Rep, 14 (2014) 492.
[103] M. Cnop, J.C. Hannaert, A. Hoorens, D.L. Eizirik, D.G. Pipeleers, Inverse
relationship between cytotoxicity of free fatty acids in pancreatic islet cells and
cellular triglyceride accumulation, Diabetes, 50 (2001) 1771-1777.
[104] H. Kaneto, Y. Kajimoto, Y. Fujitani, T. Matsuoka, K. Sakamoto, M. Matsuhisa, Y.
Yamasaki, M. Hori, Oxidative stress induces p21 expression in pancreatic islet cells:
possible implication in beta-cell dysfunction, Diabetologia, 42 (1999) 1093-1097.
[105] H. Noushmehr, E. D'Amico, L. Farilla, H. Hui, K.A. Wawrowsky, W. Mlynarski, A.
Doria, N.A. Abumrad, R. Perfetti, Fatty acid translocase (FAT/CD36) is localized on
insulin-containing granules in human pancreatic beta-cells and mediates fatty acid
effects on insulin secretion, Diabetes, 54 (2005) 472-481.
[106] S.K. Agarwal, A. Lee Burns, K.E. Sukhodolets, P.A. Kennedy, V.H. Obungu, A.B.
Hickman, M.E. Mullendore, I. Whitten, M.C. Skarulis, W.F. Simonds, C. Mateo, J.S.
Crabtree, P.C. Scacheri, Y. Ji, E.A. Novotny, L. Garrett-Beal, J.M. Ward, S.K. Libutti, H.
Richard Alexander, A. Cerrato, M.J. Parisi, A.S. Santa Anna, B. Oliver, S.C.
Chandrasekharappa, F.S. Collins, A.M. Spiegel, S.J. Marx, Molecular pathology of the
MEN1 gene, Ann N Y Acad Sci, 1014 (2004) 189-198.
[107] Y. Lu, X.Q. Fei, S.F. Yang, B.K. Xu, Y.Y. Li, Glucose-induced microRNA-17
promotes pancreatic beta cell proliferation through down-regulation of Menin, Eur
Rev Med Pharmacol Sci, 19 (2015) 624-629.
[108] Y.S. Oh, Mechanistic insights into pancreatic beta-cell mass regulation by
glucose and free fatty acids, Anat Cell Biol, 48 (2015) 16-24.
[109] R. Reynoso, L.M. Salgado, V. Calderon, High levels of palmitic acid lead to
insulin resistance due to changes in the level of phosphorylation of the insulin
receptor and insulin receptor substrate-1, Mol Cell Biochem, 246 (2003) 155-162.
[110] S.H. Um, F. Frigerio, M. Watanabe, F. Picard, M. Joaquin, M. Sticker, S.
Fumagalli, P.R. Allegrini, S.C. Kozma, J. Auwerx, G. Thomas, Absence of S6K1 protects
against age- and diet-induced obesity while enhancing insulin sensitivity, Nature,
431 (2004) 200-205.
[111] L. Khamzina, A. Veilleux, S. Bergeron, A. Marette, Increased activation of the
mammalian target of rapamycin pathway in liver and skeletal muscle of obese rats:
possible involvement in obesity-linked insulin resistance, Endocrinology, 146
(2005) 1473-1481.
[112] J.M. Bard, M.A. Charles, I. Juhan-Vague, P. Vague, P. Andre, M. Safar, J.C.
Fruchart, E. Eschwege, B.S. Group, Accumulation of triglyceride-rich lipoprotein in
subjects with abdominal obesity: the biguanides and the prevention of the risk of
obesity (BIGPRO) 1 study, Arterioscler Thromb Vasc Biol, 21 (2001) 407-414.
99
[113] D.C. Chan, G.F. Watts, P.H. Barrett, J.C. Mamo, T.G. Redgrave, Markers of
triglyceride-rich lipoprotein remnant metabolism in visceral obesity, Clin Chem, 48
(2002) 278-283.
[114] G.F. Lewis, A. Carpentier, K. Adeli, A. Giacca, Disordered fat storage and
mobilization in the pathogenesis of insulin resistance and type 2 diabetes, Endocr
Rev, 23 (2002) 201-229.
[115] Y.Z. Liu, X. Cheng, T. Zhang, S. Lee, J. Yamauchi, X. Xiao, G. Gittes, S. Qu, C.L.
Jiang, H.H. Dong, Effect of Hypertriglyceridemia on Beta Cell Mass and Function in
ApoC3 Transgenic Mice, J Biol Chem, 291 (2016) 14695-14705.
[116] T.L. Halvorsen, G.M. Beattie, A.D. Lopez, A. Hayek, F. Levine, Accelerated
telomere shortening and senescence in human pancreatic islet cells stimulated to
divide in vitro, J Endocrinol, 166 (2000) 103-109.
[117] Q. Chen, B.N. Ames, Senescence-like growth arrest induced by hydrogen
peroxide in human diploid fibroblast F65 cells, Proc Natl Acad Sci U S A, 91 (1994)
4130-4134.
[118] H. Iwasa, J. Han, F. Ishikawa, Mitogen-activated protein kinase p38 defines the
common senescence-signalling pathway, Genes Cells, 8 (2003) 131-144.
[119] G. Kroemer, G. Marino, B. Levine, Autophagy and the integrated stress
response, Mol Cell, 40 (2010) 280-293.
[120] E. Kim, P. Goraksha-Hicks, L. Li, T.P. Neufeld, K.L. Guan, Regulation of TORC1
by Rag GTPases in nutrient response, Nat Cell Biol, 10 (2008) 935-945.
[121] Y. Sancak, T.R. Peterson, Y.D. Shaul, R.A. Lindquist, C.C. Thoreen, L. Bar-Peled,
D.M. Sabatini, The Rag GTPases bind raptor and mediate amino acid signaling to
mTORC1, Science, 320 (2008) 1496-1501.
[122] Y. Sancak, L. Bar-Peled, R. Zoncu, A.L. Markhard, S. Nada, D.M. Sabatini,
Ragulator-Rag complex targets mTORC1 to the lysosomal surface and is necessary
for its activation by amino acids, Cell, 141 (2010) 290-303.
[123] L.M. Dickson, M.K. Lingohr, J. McCuaig, S.R. Hugl, L. Snow, B.B. Kahn, M.G.
Myers, Jr., C.J. Rhodes, Differential activation of protein kinase B and p70(S6)K by
glucose and insulin-like growth factor 1 in pancreatic beta-cells (INS-1), J Biol Chem,
276 (2001) 21110-21120.
[124] D. Menon, D. Salloum, E. Bernfeld, E. Gorodetsky, A. Akselrod, M.A. Frias, J.
Sudderth, P.H. Chen, R. DeBerardinis, D.A. Foster, Lipid Sensing by mTOR via de
novo Synthesis of Phosphatidic Acid, J Biol Chem, (2017).
[125] Y. Fang, M. Vilella-Bach, R. Bachmann, A. Flanigan, J. Chen, Phosphatidic acid-
mediated mitogenic activation of mTOR signaling, Science, 294 (2001) 1942-1945.
[126] A. Toschi, E. Lee, L. Xu, A. Garcia, N. Gadir, D.A. Foster, Regulation of mTORC1
and mTORC2 complex assembly by phosphatidic acid: competition with rapamycin,
Mol Cell Biol, 29 (2009) 1411-1420.
[127] M.K. Lingohr, L.M. Dickson, J.F. McCuaig, S.R. Hugl, D.R. Twardzik, C.J. Rhodes,
Activation of IRS-2-mediated signal transduction by IGF-1, but not TGF-alpha or
EGF, augments pancreatic beta-cell proliferation, Diabetes, 51 (2002) 966-976.
[128] M.K. Lingohr, L.M. Dickson, C.E. Wrede, I. Briaud, J.F. McCuaig, M.G. Myers, Jr.,
C.J. Rhodes, Decreasing IRS-2 expression in pancreatic beta-cells (INS-1) promotes
apoptosis, which can be compensated for by introduction of IRS-4 expression, Mol
Cell Endocrinol, 209 (2003) 17-31.
100
[129] D.J. Withers, D.J. Burks, H.H. Towery, S.L. Altamuro, C.L. Flint, M.F. White, Irs-2
coordinates Igf-1 receptor-mediated beta-cell development and peripheral insulin
signalling, Nat Genet, 23 (1999) 32-40.
[130] D.J. Withers, J.S. Gutierrez, H. Towery, D.J. Burks, J.M. Ren, S. Previs, Y. Zhang,
D. Bernal, S. Pons, G.I. Shulman, S. Bonner-Weir, M.F. White, Disruption of IRS-2
causes type 2 diabetes in mice, Nature, 391 (1998) 900-904.
[131] I. Briaud, L.M. Dickson, M.K. Lingohr, J.F. McCuaig, J.C. Lawrence, C.J. Rhodes,
Insulin receptor substrate-2 proteasomal degradation mediated by a mammalian
target of rapamycin (mTOR)-induced negative feedback down-regulates protein
kinase B-mediated signaling pathway in beta-cells, J Biol Chem, 280 (2005) 2282-
2293.
[132] T. Yuan, S. Rafizadeh, K.D. Gorrepati, B. Lupse, J. Oberholzer, K. Maedler, A.
Ardestani, Reciprocal regulation of mTOR complexes in pancreatic islets from
humans with type 2 diabetes, Diabetologia, 60 (2017) 668-678.
[133] K.T. Yang, J.A. Bayan, N. Zeng, R. Aggarwal, L. He, Z. Peng, A. Kassa, M. Kim, Z.
Luo, Z. Shi, V. Medina, K. Boddupally, B.L. Stiles, Adult-onset deletion of Pten
increases islet mass and beta cell proliferation in mice, Diabetologia, 57 (2014) 352-
361.
[134] B.L. Stiles, C. Kuralwalla-Martinez, W. Guo, C. Gregorian, Y. Wang, J. Tian, M.A.
Magnuson, H. Wu, Selective deletion of Pten in pancreatic beta cells leads to
increased islet mass and resistance to STZ-induced diabetes, Mol Cell Biol, 26 (2006)
2772-2781.
[135] C.B. Rountree, W. Ding, L. He, B. Stiles, Expansion of CD133-expressing liver
cancer stem cells in liver-specific phosphatase and tensin homolog deleted on
chromosome 10-deleted mice, Stem Cells, 27 (2009) 290-299.

Abstract (if available)

Abstract Hyperlipidemia is one of the hallmarks of Diabetes. β cells are exposed to high levels of free fatty acids (FFAs) along with glucose during Diabetes development. In this study, we focused on the exposure-dependent effect of FFAs on β cells by using a High Fat Diet (HFD) model in mice and Palmitic Acid (PA) treatment in INS-1 cell line. FFAs have been earlier observed to have pro-proliferative effect upon short-term exposure and can lead to cell death upon long-term exposure in β cells. However, the molecular players modulating such effect are still unknown. Since β cell proliferation is regulated by cell cycle proteins such as Cyclins, CDKs (Cyclin dependent kinases) and CDKIs (Cyclin dependent kinase inhibitors), we hypothesized that short term exposure of FFAs will increase cell proliferation while the long term will decrease cell growth via the cell cycle regulatory proteins. To test this hypothesis, we fed mice with HFD for 14 days and 2 months (short-term) and 9 & 14 months (long-term). Indeed, β cell proliferation was significantly upregulated after short-term exposure, which was accompanied by both hyperglycemia and hyperlipidemia. This β cell proliferation was due to up-regulation of Cyclin D2 in mice islets. Our in vitro PA treatment results further demonstrated that Cyclin D2 induction was regulated by the lipid mediated activation of mTOR pathway. RNA seq analysis of the islets from HFD vs NC fed mice supported significant enrichment of the genes involved in the mTOR pathway. Additionally, this result was confirmed in mice by using rapamycin (mTOR inhibitor) which abolished HFD mediated proliferative effect. ❧ Furthermore, long term HFD exposure decreased β cell proliferation. Diabetic individuals experience decrease in β cell mass due to which they can no longer maintain glucose homeostasis. In our long term HFD treatment model, plasma triglycerides (TG) were significantly upregulated, along with decrease in Cyclin D2 and increase in p27 levels in islets. In vitro treatment of INS-1 cells with TG confirmed its effect on both Cyclin D2 and p27, suggesting a direct impact of lipids on β cell proliferation and possible effect of TG on β cell senescence. ❧ Together, we found that FFAs regulate β cell proliferation in an exposure dependent manner via mTOR, Cyclin D2 and p27. Studies identifying molecular regulators of β cell proliferation can contribute in targeted therapies that can increase β cell mass in Diabetic individuals and alleviate Diabetes symptoms.

Linked assets

University of Southern California Dissertations and Theses

Conceptually similar

PDF

Protein kinase Bα (AKT1) but not protein kinase Bβ (AKT2) controls pancreatic β cell growth and survival

PDF

Akt1 deletion decrease proliferation in aged pancreatic beta-cells by arresting cell cycle in S phase

PDF

Steatosis induces Wnt/β-catenin pathway to stimulate proliferation of hepatic tumor initiating cells and promote liver cancer development

PDF

Pten regulates beta-cell regeneration intrinsically and independently of development

PDF

PTEN loss antagonizes aging through promoting regeneration and prevents oxidative stress induced cell death

PDF

The overlap between mTOR signaling, rapamycin and cellular senescence

Asset Metadata

Creator Aggarwal, Richa (author)

Core Title Short term high fat diet (HFD) stimulates β cell proliferation through mTOR while the prolonged treatment induces β cell senescence via p27

School School of Pharmacy

Degree Doctor of Philosophy

Degree Program Molecular Pharmacology and Toxicology

Publication Date 04/28/2018

Defense Date 03/08/2017

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag beta cell mass,beta cell proliferation,Diabetes,high fat diet,mTOR,OAI-PMH Harvest,obesity

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Stiles, Bangyan (committee chair), Okamoto, Curtis (committee member), Watanabe, Richard (committee member)

Creator Email richa.1009@gmail.com,richaagg@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-366719

Unique identifier UC11258020

Identifier etd-AggarwalRi-5293.pdf (filename),usctheses-c40-366719 (legacy record id)

Legacy Identifier etd-AggarwalRi-5293.pdf

Dmrecord 366719

Document Type Dissertation

Rights Aggarwal, Richa

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA