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Mechanisms of Glucagon-Like Peptide-2-Mediated Effects on
Intestinal Barrier Function in Health and Irinotecan-Induced
Enteritis
by
Charlotte Xiaoman Dong
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Graduate Department of Physiology
University of Toronto
© Copyright by Charlotte Xiaoman Dong (2013)
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Mechanisms of Glucagon-Like Peptide-2-Mediated Effects on
Intestinal Barrier Function in Health and Irinotecan-Induced Enteritis
Charlotte X. Dong
Master of Science
Department of Physiology
University of Toronto
2013
ABSTRACT
Glucagon-like peptide-2 (GLP-2) is an intestinal hormone that promotes gut growth through
an insulin-like growth factor (IGF)-1 and intestinal epithelial (IE)-IGF-1 receptor (R)-
dependent pathway. GLP-2 also promotes epithelial barrier function by as yet unknown
mechanisms. I hypothesized that GLP-2-mediated effects on barrier function requires the IE-
IGF-1R. Chronic GLP-2 treatment enhanced barrier function by decreasing gastrointestinal
permeability in vivo and increasing jejunal resistance ex vivo. These responses were
abolished in inducible IE-IGF-1R knockout (KO) animals. Additionally, epithelial sealing
tight junctional proteins claudin-3 and -7 were upregulated by GLP-2 in control but not KO
mice. Moreover, IE-IGF-1R deletion induced a shift in occludin localization from apical to
intracellular domains. In contrast, in irinotecan-induced enteritis, GLP-2 normalized
epithelial barrier function in control animals, but continued to be ineffective in KO mice.
Collectively, the effects of GLP-2 on barrier function are dependent on the IE-IGF-1R and
involve modulation of the tight junctional complex.
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ACKNOWLEDGEMENTS
Graduate school has been a large part of my life in the past two years, and my M.Sc.
experience has catalyzed the growth of not only my research abilities, but also personal
skills. There are many people to thank who have been encouraging and instrumental in this
M.Sc. journey. First and foremost, I would like to thank my extraordinary supervisor and
long-time mentor, Dr. Pat Brubaker. Her enthusiasm in science is contagious and her
dedication to teaching motivates me to continuously strive for excellence. At times of
success, I was congratulated with joy while still reminded of areas for improvement, and at
times of failure, I was forgiven with never-ceasing support and understanding. I would like to
extend my thanks to my committee members – Dr. Nicola Jones, for the excellent and
thought-provoking questions during committee meetings, Dr. Tanja Gonska, for providing
expertise and on-going guidance on the electrophysiology studies, and Dr. Tony Lam for
providing an exceptional seminar course that has expanded my knowledge. I am grateful for
all your invaluable advice and feedback throughout this project.
I also want to thank all the past and present members the Brubaker Lab for help and
companionship. I am especially grateful to Wen Zhao and Chloe Solomon for all your hard
work and contributions, especially in conducting the immunofluorescence experiments and
the beautiful micrographs. I am indebted to Dr. Katie Rowland for the tremendous amount of
effort invested into establishing the IE-IGF-1R KO mouse model, and to Monika Poreba,
who taught me the fundamentals of scientific research when I was an undergraduate student.
Graduate school was a unique experience enhanced and enriched by fellow graduate
students, Jasleen Chahal, Samantha Li and Kaori Yamada, and I was fortunate to have the
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knowledgeable post-doctoral fellows, Manuel Gil Lozano and Jeffrey Gagnon, in the lab for
advice and suggestions.
I would also like to thank our collaborators Dr. Cameron Ackerley for the electron
microscopy studies, Dr. Sylvie Robine, Dr. Martin Holzenberger and Dr. Rohit Kulkarni for
the transgenic animals. A special thank you goes to Wan Ip for teaching me and
troubleshooting with me during Ussing chamber studies. Many thanks to our animal facility
staff Dr. Kate Banks, Diana Hiesl, Leila Tick, Tracy McCook, Sara Johnson, Nancy Thomas
and Mike Grant for expertise and transportation of animals. I also appreciate the awesome
physiology administrative team, Rosalie, Eva and Colleen for always being extremely
reliable, efficient and helpful, and the Department of Physiology for numerous opportunities
in academic development.
Lastly, I want to thank my parents and Shawn Yin for love and support. Graduate
school has transformed me into the person that I am today, and I will carry the invaluable
friendships and lessons with me into future endeavours.
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TABLE OF CONTENTS
ACKNOWLEDGEMENTS ........................................................................................................... iii
TABLE OF CONTENTS ................................................................................................................ v
LIST OF FIGURES ...................................................................................................................... vii
Dissemination of Work Arising from this Thesis .......................................................................... ix
LIST OF ABBREVIATIONS ......................................................................................................... x
1 INTRODUCTION...................................................................................................................... 1
1.1 Rationale ............................................................................................................................. 1
1.2 Glucagon-like peptide-2 ...................................................................................................... 2
1.2.1 Discovery ................................................................................................................ 2
1.2.2 Gene expression ...................................................................................................... 3
1.2.3 Synthesis ................................................................................................................. 3
1.2.4 Secretion ................................................................................................................. 6
1.2.5 Metabolism and clearance ..................................................................................... 10
1.2.6 The GLP-2R .......................................................................................................... 10
1.3 GLP-2 and the gastrointestinal tract ................................................................................. 12
1.3.1 Actions of endogenous GLP-2 .............................................................................. 12
1.3.2 Actions of exogenous GLP-2 ................................................................................ 14
1.4 Insulin-like growth factors ................................................................................................ 17
1.4.1 IGF peptides .......................................................................................................... 18
1.4.2 Metabolism and clearance ..................................................................................... 19
1.4.3 The IGF-1R ........................................................................................................... 19
1.5 Epithelial barrier function ................................................................................................. 24
1.6 Hypothesis and specific aims ............................................................................................ 28
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2 MATERIALS AND METHODS ............................................................................................. 30
2.1 Animals ............................................................................................................................. 30
2.2 Intestinal permeability ...................................................................................................... 34
2.3 Microscopy ....................................................................................................................... 35
2.4 Immunoblotting ................................................................................................................. 36
2.5 Statistical analyses ............................................................................................................ 37
3 RESULTS ................................................................................................................................ 38
3.1 Validation of the IE-IGF-1R KO mouse model ................................................................ 38
3.2 The IE-IGF-1R was essential for GLP-2-enhanced barrier function ................................ 40
3.3 No change in tight junctional complex ultrastructure was detected by electron
microscopy ........................................................................................................................ 40
3.4 GLP-2 upregulated claudin-3 and -7 expression via the IE-IGF-1R ................................ 44
3.5 Deletion of the IE-IGF-1R altered occludin localization .................................................. 48
3.6 Scaffolding ZO-1 was unchanged with IE-IGF-1R deletion and/or GLP-2
treatment ........................................................................................................................... 48
3.7 The IE-IGF-1R was required for GLP-2-normalized barrier function in irinotecan-
induced enteritis ................................................................................................................ 51
3.8 GLP-2-normalized barrier function may require mechanisms independent of the
tight junctions .................................................................................................................... 55
4 DISCUSSION .......................................................................................................................... 60
5 APPENDIX .............................................................................................................................. 68
5.1 Intestinal permeability and diabetes mellitus .................................................................... 68
5.2 Material and Methods ....................................................................................................... 68
5.3 Summary of STZ studies .................................................................................................. 69
6 REFERENCES ......................................................................................................................... 72
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LIST OF FIGURES
Figure 1.1 Tissue-specific post-translational processing of proglucagon. ................................ 5
Figure 1.2 Secretogogues of the enteroendocrine L cell. .......................................................... 8
Figure 1.3 Structure and intracellular signaling of the IGF-1R. ............................................. 21
Figure 1.4 Intestinal epithelial apical junction complexes. ..................................................... 25
Figure 2.1 Generation of the IE-IGF-1R KO mouse using the Cre/lox system ...................... 31
Figure 2.2 Male mice appeared more sensitive to irinotecan than female mice. .................... 33
Figure 3.1 Identification and validation of the IE-IGF-1R KO mouse model. ....................... 39
Figure 3.2 GLP-2-enhanced barrier function was reduced in KO mice. ................................ 41
Figure 3.3 Ultrastructure of tight junctional proteins was unchanged. ................................... 43
Figure 3.4 Protein expression levels of tight junctional claudins. .......................................... 45
Figure 3.5 Immunofluorescence of claudin-3, -7 and -15. ..................................................... 47
Figure 3.6 Protein expression levels of occludin and ZO-1. ................................................... 49
Figure 3.7 Subcellular localization of occludin and ZO-1. ..................................................... 50
Figure 3.8 GLP-2-restored crypt-villus growth in irinotecan-induced enteritis required the IE-
IGF-1R. ................................................................................................................................... 53
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Figure 3.9 GLP-2-normalized barrier function required the IE-IGF-1R in irinotecan-induced
enteritis .................................................................................................................................... 54
Figure 3.10 Protein expression of tight junctions in irinotecan-induced enteritis. ................. 57
Figure 3.11 Localization of tight junctions in irinotecan-induced enteritis. ........................... 59
Figure 5.1 STZ did not induce diabetes nor impaired barrier function .................................. 71
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Dissemination of Work Arising from this Thesis
Contents from chapters 3 & 4 were submitted for publication/published as:
Dong,C.X., W.Zhao, C.Solomon, K.J.Rowland, C.Ackerley, S.Robine,
M.Holzenberger, T.Gonska and P.L.Brubaker. Glucagon-like peptide-2 effects on the
murine gut barrier require the epithelial insulin-like growth factor-1 receptor.
Submitted, 2013.
Dong,C.X., C.Solomon, W.Zhao, T.Gonska and P.L.Brubaker. Role of the intestinal
epithelial-insulin-like growth factor-1 receptor in glucagon-like peptide-2-mediated
enhancement of intestinal barrier function. Dig Dis Week, 697-OR, 2013.
Additional publication arising during the course of my graduate studies:
Dong,C.X. and P.L.Brubaker. Ghrelin, the proglucagon-derived peptides and peptide
YY in nutrient homeostasis. Nat Rev Gastroenterol Hepatol, 9:705-715, 2012
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LIST OF ABBREVIATIONS
ANOVA Analysis of variance
bp Base pair
cAMP Cyclic adenosine monophosphate
Cre Cyclization recombination
CREB cAMP response element-binding
DAPI 4'-6-diamidino-2-phenylindole
DNA Deoxyribonucleic acid
DPP-4 Dipeptidyl peptidase-4
EGF Epidermal growth factor
ER Estrogen receptor
eNOS Endothelial nitric oxide synthase
ERK Extracellular signal-regulated kinase
FD4 Fluorescein isothiocyanate dextran 4000
GI Gastrointestinal
GLP-2 Glucagon-like peptide-2
GLP-2R Glucagon-like peptide-2 receptor
GLUT-2 Glucose transporter-2
GPR G-protein coupled receptor
H&E Hematoxylin & eosin
IE Intestinal epithelial
IGF Insulin-like growth factor
IGF-1R Insulin-like growth factor-1 receptor
IGF-2R Insulin-like growth factor-2 receptor
IGFBP Insullin-like growth factor binding protein
IP Intervening peptide
IR Insulin receptor
IRS Insulin receptor substrate
JAM Junctional adhesion molecule
KGF Keratinocyte growth factor
KO Knockout
L1 Leucine-rich repeat domain 1
L2 Leucine-rich repeat domain 2
LI Large intestine
MAPK Mitogen-activated protein kinase
MEK MAPK/ERK kinase
mRNA Messenger ribonucleic acid
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PBS Phosphate-buffered saline
PC Prohormone convertase
PCR Polymerase chain reaction
PDK-1 3-phosphoinositide dependent protein
kinase-1
PI3K Phosphoinositide-3-kinase
PKA Protein kinase A
SEMF Subepithelial myofibroblast
SGLT-1 Sodium-dependent glucose cotransporter-1
SBS Short bowel syndrome
SI Small intestine
TBST Tris-buffered saline with Tween
TGF
VEGF
Transforming growth factor
Vascular endothelial cell growth factor
VIP Vasoactive intestinal polypeptide
ZO Zona occluden
Symbols and units
~ Approximately
% Percent
°C Degrees Celsius
g Grams
hr Hours
kDa Kilodaltons
l Litres
M Molar (moles/litre)
p Statistical p-value
s Seconds
SEM Standard error of the mean
wk Weeks
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1 INTRODUCTION
1.1 Rationale
Glucagon-like peptide-2 (GLP-2) is a potent nutrient-dependent intestinal growth factor that
promotes intestinal growth and function (Dong and Brubaker, 2012; Dube and Brubaker,
2007; Rowland and Brubaker, 2011). Teduglutide, a long acting analogue of GLP-2, has
recently been approved for the treatment of adult Short Bowel Syndrome (SBS). Teduglutide
is also in phase 2 clinical trials for Crohn’s disease (NCT00072839; NCT00308438) and is in
preclinical studies for pediatric SBS (npsp.com). Because of such clinical advances related to
the application of GLP-2, it is crucial to delineate its mechanism(s) of action. In particular, it
remains unknown how GLP-2 improves intestinal barrier function. Based on GLP-2 receptor
(R) distribution, it is known that GLP-2 does not directly act upon its target intestinal
epithelial cells but, rather, via indirect pathways that activate different downstream mediators
to result in corresponding effects (Guan et al., 2006; Orskov et al., 2005; Yusta et al., 2000).
We have previously demonstrated that GLP-2 can signal through another growth factor,
insulin-like growth factor (IGF)-1 (Dube et al., 2006; Dube et al., 2008), acting on its
receptor localized to the intestinal epithelium (IE-IGF-1R) to promote mucosal growth
(Rowland et al., 2011). Since no GLP-2 mediator has been identified to date that is
responsible for its effects on enhancing epithelial barrier function, the goal of this study was
to determine the requirement for the IE-IGF-1R in this pathway and the mechanisms
involved.
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1.2 Glucagon-like peptide-2
1.2.1 Discovery
In 1961, Unger et al. published a study on generation of novel glucagon antisera that
also reported serendipitous identification of glucagon-like immunoreactive peptides released
by the gastrointestinal (GI) tract (Unger et al., 1961). The glucagon-like peptides were
designated enteroglucagon and are now known to be co-secreted with the related peptides,
GLP-1 and GLP-2. However, GLP-2 function remained elusive for many years. Interestingly,
a rare renal glucagonoma-bearing patient was reported to have small bowel hyperplasia
(Gleeson et al., 1971). The abnormal intestinal growth regressed following nephrectomy,
which shed light on the potential existence of a tumour product with intestinotrophic
properties. A decade later, two more glucagonoma case studies reported similar phenotypes
(Jones et al., 1983; Stevens et al., 1984). Meanwhile, cloning of preproglucagon identified, in
addition to glucagon, GLP-1 and GLP-2 to be tandemly encoded in a single gene (Bell et al.,
1983a; Bell et al., 1983b; Lund et al., 1982). Finally, with advances in the knowledge of
GLP-2 structure and generation of its synthetic forms, GLP-2 was determined to be the
glucagonoma secretory product that caused profound gut growth (Drucker et al., 1996).
Thenceforth, a large number of studies have expanded our knowledge on multiple intestinal
specific actions of GLP-2, including the enhancement of barrier function (Benjamin et al.,
2000).
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1.2.2 Gene expression
The major source of GLP-2 comes from the intestinal L cell where proglucagon gene
expression is stimulated by feeding and amplified by refeeding after fasting (Hoyt et al.,
1996). Fibre and short-chain fatty acids are particularly potent in promoting gene expression
(Reimer and McBurney, 1996; Tappenden et al., 1996). Insulin is also elevated in response to
feeding and increases proglucagon mRNA through Wnt-signaling (Chen et al., 1989). In
addition, in vitro upregulators of proglucagon gene expression include cAMP and the
transcription factors Pax6 and TCF4 (Drucker and Brubaker, 1989; Hill et al., 1999; Yi et al.,
2005).
1.2.3 Synthesis
The preproglucagon gene is located on chromosome 2 in mammals and is comprised
of 6 exons (Bell et al., 1983a; Schroeder et al., 1984). The major translated peptides, namely
glucagon, GLP-1 and GLP-2, are encoded by different exons, suggestive of evolutionary
internal triplication of a common ancestral glucagon gene (Bell et al., 1983a). GLP-1 and
GLP-2 sequences are highly conserved across mammalian species; human GLP-2 differs
from that of rat and mouse by 1 and 2 amino acids, respectively, whereas GLP-1 is 100%
conserved (Mojsov et al., 1986; White and Saunders, 1986).
Proglucagon is a 160-amino acid protein expressed in the pancreatic α cell, the
intestinal L cell, and, to a lesser extent, the brain, which undergoes differential post-
translational processing in a tissue-specific manner (Figure 1.1) (Mojsov et al., 1986).
Proteolytic cleavage of proglucagon in the L cell is mediated by prohormone convertase (PC)
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1/3 to liberate the 33-amino acid GLP-2 along with GLP-1 and enteroglucagon (also known
as glicentin or oxyntomodulin, a C-terminal extended form of glucagon) (Dhanvantari et al.,
1996; Rouille et al., 1995; Rothenberg et al., 1996). The importance of PC1/3 in producing
GLP-2 has been demonstrated by reductions in GLP-2 levels in PC1/3 null mice and patients
with PC1/3 mutations (Jackson et. al, 2003; Zhu et al., 2002). Alternatively, proglucagon in
the pancreatic α cell is processed by PC2 to release glucagon, glicentin-related pancreatic
peptide and the major proglucagon fragment, but not GLP-2 (Rouille et al., 1994).
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Figure 1.1 Tissue-specific post-translational processing of proglucagon.
The 160-amino acid precursor peptide, proglucagon, is cleaved by prohormone convertase
(PC) 1/3 in the intestinal L cell and likely, the brain, to release GLP-2 along with GLP-1 and
oxyntomodulin or glicentin. Alternatively, proglucagon is cleaved by PC2 in the pancreatic
α cell to liberate glucagon, glicentin-related pancreatic peptide (GRPP) and major
proglucagon fragment (MPGF). IP-1/2, intervening peptide-1/2.
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1.2.4 Secretion
The GLP-2-producing enteroendocrine L cell is flask-shaped and open-type, with
microvilli that contact the intestinal lumen and endocrine granules residing near the basal
lamina, in proximity with neural and vascular structures (Eissele et al., 1992; Larsson et al.,
1975). This architectural layout indicates that L cells can synthesize and secrete their
granular content in response to luminal, neural and circulatory inputs. Along the aboral axis,
L cells are more densely distributed along the distal gut, but can also be found in the upper
GI tract in markedly lower numbers (Eissele et al., 1992; Larsson et al., 1975). Interestingly,
proximal and distal L cells are not homogeneous and differ by gene expression profiles and
responsivity to secretogogues (Habib et al., 2012; Geraedts et al., 2012; Reimann et al., 2008;
Egerod et al., 2012).
GLP-2 is secreted in response to nutrient ingestion, which increases plasma GLP-2
levels from 15-20 pmol/L under fasting conditions to 30-60 pmol/L in humans (Xiao et al.,
1999; Brubaker et al., 1997a). Secretion of GLP-2 exhibits a biphasic pattern with an acute
rapid increase followed by a delayed prolonged response (Orskov et al., 1986; Xiao et al.,
1999). Specifically, nutrients in the duodenum acutely stimulate the vagus nerve to indirectly
activate ileal and colonic L cells via muscarinic receptors (Anini and Brubaker, 2003a; Rocca
and Brubaker, 1999; Xiao et al., 1999), whereas the delayed response occurs following
transit of luminal nutrients distally to directly stimulate L cells (Figure 1.2) (Iakoubov et al.,
2007). Luminal nutrient-triggered GLP-2 release is predominantly mediated by various forms
of short- and long-chain fatty acids (Iakoubov et al., 2007; Iakoubov et al., 2009; Rocca and
Brubaker, 1999; Hirasawa et al., 2005; Edfalk et al., 2008; Poreba et al., 2012). Glucose and
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bile acids also stimulate secretion, although the physiological significance of glucose in the
distal gut remains to be clarified (Parker et al., 2012; Gribble et al., 2003; Thomas et al.,
2009). In addition, L cell secretagogues include hormones and neuropeptides such as
glucose-dependent insulinotrophic polypeptide, gastrin-releasing peptide, insulin and leptin
(Anini and Brubaker, 2003b; Lim et al., 2009; Roberge and Brubaker, 1993; Roberge et al.,
1996; Chisholm and Greenberg, 2002).
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Figure 1.2 Secretogogues of the enteroendocrine L cell.
Vagal stimulation occurs via acetylcholine (ACh) to activate the M1 muscarinic receptor
(M1R). Insulin and leptin acts through canonical pathways (IR, insulin receptor; PI3K,
phosphatidylinositol-3 kinase-1; MEK, MAPK/ERK kinase; LR, leptin receptor; STAT3,
signal transducer activator of transcription 3). Fatty acids (FAs) and their derivatives (OEA,
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oleoylethanoloamide; 2-oleoylglycerol) signal through G protein-coupled receptors (GPRs)
followed by PKA or PKC activation, whereas uptake of oleic acid (OA) via fatty acid
transporter (FATP) 4 is followed by PKCζ activation. Glucose is transported by sodium
glucose cotransporter (SGLT) 1 to cause depolarization (ψ)-induced calcium (Ca2+
) flux.
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1.2.5 Metabolism and clearance
Upon secretion, GLP-2 has a short half-life of ~7 minutes in rats and humans
rendered by extensive renal clearance at the rate of glomerular filtration and rapid enzymatic
degradation by circulating dipeptidyl peptidase-4 (DPP-4) (Ruiz-Grande et al., 1990; Tavares
et al., 2000). Proteolytic DPP-4 cleaves GLP-2 at the N-terminal penultimate Ala2, and
exhibits higher activity in rats than in mice (Tavares et al., 2000; Hartmann et al., 2000). The
cleavage product GLP-23-33
is also cleared by the renal system. Importantly, the cleavage
product GLP-23-33
remains active and agonistic activities can be detected at high
concentrations (Drucker et al., 1997a; Shin et al., 2005; Hartmann et al., 2000; Tavares et al.,
2000). Thus, most studies on the actions of GLP-2 in vivo, including those in the present
study, use the DPP-4-resistant long-acting analogue, human Gly2-GLP-2 or teduglutide, that
contains a substitution of Ala2 with the DPP-4-resistant amino acid Gly
2 (Tavares et al.,
2000; Drucker et al., 1997b). This GLP-2 analog has been demonstrated to bind to the murine
GLP-2R (Shin et al., 2005), and is functional in all rodent models studied to date.
1.2.6 The GLP-2R
The GLP-2R is a member of the class B glucagon receptor family of GPRs with
typical heptahelical typology, and is characterized by a long N-terminal extracellular tail
containing two loops, which is important for ligand binding (Mayo et al., 2003). Cloning of
the GLP-2R in humans revealed that the receptor is encoded within chromosome 17 and
shares 81.6% amino acid sequence homology with the rat GLP-2R (Munroe et al., 1999),
while the murine GLP-2R is 85-90% identifical to the human and rat receptors (Shin et al.,
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2005). Furthermore, the human and rat GLP-2R exhibit the same profile of peptide-binding
specificity (Munroe et al., 1999). To date, no GLP-2R or GLP-2 mutation has been identified
in humans.
Localization of the GLP-2R is highly restricted to the GI tract, with very limited
expression also found in the hypothalamus, lungs and cervix (Koehler et al., 2005; Munroe et
al., 1999; Yusta et al., 2000). In the GI tract, GLP-2R density is higher within the SI, and is
particularly abundant along the jejunal segment (Munroe et al., 1999; Yusta et al., 2000).
Interestingly, the GLP-2R is not present on the crypt or villus epithelial cells that ultimately
exert most of the effects of GLP-2 (Yusta et al., 2000). Rather, GLP-2R has been detected in
the subepithelial myofibroblasts (SEMFs), rare enteroendocrine cells and the enteric nerve
plexus (Bjerknes and Cheng, 2001; Guan et al., 2006; Orskov et al., 2005). It was therefore
postulated that the effects of GLP-2 are transduced indirectly via downstream mediators
secreted from GLP-2R-expressing cells (Yusta et al., 2000). Evidence supporting this notion
has accumulated to identify several mediators for multiple aspects of GLP-2 actions, which
are addressed individually in section 1.3.2.
Pathway mapping of GLP-2R intracellular signaling has been rendered difficult by a
lack of appropriate cell models. In vitro studies on GLP-2R signaling are often done by
transfecting the GLP-2R into heterologous cell models, including BHK-, COS- and DLD-1-
GLP-2R transfected cells (Munroe et al., 1999; Thulesen et al., 2002; Estall et al., 2004;
Yusta et al., 1999; Yusta et al., 2000). These cell lines respond to GLP-2 stimulation with
increased levels of intracellular cAMP. Similarly, a GLP-2R-induced cAMP response has
also been demonstrated in primary cultures of rat intestinal mucosa (Walsh et al., 2003), fetal
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rat intestine (Dube et al., 2006), astrocytes (Velazquez et al., 2003), hippocampal cells
(Lovshin et al., 2004) and mouse muscle strips (Anini et al., 2007). Furthermore, among the
cell models available that naturally express the GLP-2R, the human intestinal epithelial FHC
cell line also exhibits the cAMP response (Sams et al., 2006). Downstream of cAMP, shown
in BHK-GLP-2R transfected fibroblasts, PKA signaling stimulates CREB and AP-1 (Yusta et
al., 1999). In contrast, a cAMP response is not detected in primary cultures of rodent SEMFs
or enteric neurons, or in the human HeLa cell line – these cell types naturally express the
GLP-2R (Koehler et al., 2005; Leen et al., 2011; de Heuval et al., 2012). Rather, in the
SEMFs and enteric neurons, the PI3K/AKT pathway is activated (Leen et al., 2011). This
leads to increased IGF-1 mRNA transcript levels in the SEMF cells. A GLP-2R-activated
PI3K pathway has also been reported in BHK-GLP-2R transfected cells (Yusta et al., 2002).
Furthermore, a recent study shows that GLP-2 signalling in hippocampal neurons requires
PI3K-dependent Akt activation (Shi et al., 2013).
1.3 GLP-2 and the gastrointestinal tract
1.3.1 Actions of endogenous GLP-2
Cumulative data on manipulation of GLP-2 actions suggest that endogenously
produced GLP-2 exerts modest trophic effects. Three experimental approaches have been
used in the past, each of which is subject to limitations. Initial experiments used the
immunoneutralization technique targeting circulating GLP-2 and revealed reduced adaptive
intestinal growth in a disease model of type 1 diabetes (Hartmann et al., 2000). However,
immunoneutralization may only achieve partial inhibition of circulating GLP-2 and is
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therefore not feasible for chronic studies due to a possible compensatory increase in GLP-2
secretion. Inhibition of the GLP-2R using the truncated GLP-23-33
at antagonistic doses is an
alternative pharmacologic technique. GLP-23-33
reduces mucosal growth under basal
conditions, as well as upon refeeding after fasting for 24 hours and following 4 weeks of
administration (Iakoubov et al., 2009; Shin et al., 2005; Nelson et al., 2008).
Notwithstanding, GLP-23-33
can also exhibit partial agonistic activities if not administered at
appropriate doses, thereby yield confounding results (Thulesen et al., 2002). Genetic
manipulation of the GLP-2R is another approach to investigate the actions of endogenous
GLP-2. Interestingly, GLP-2R KO mice do not exhibit abnormal intestinal morphology or
growth impairment (Lee et al., 2012; Bahrami et al., 2010). Under healthy conditions, the
largely normal phenotype of the KO mice with GLP-2R deleted embryonically is suggested
to be due to compensatory effects throughout development. Nonetheless, under fasting
conditions, refeeding-induced intestinal growth is absent in the GLP-2R KO mice, thereby
indicating a requirement for endogenous GLP-2 (Bahrami et al., 2010). Furthermore, the
growth response observed in refed mice is dependent upon GLP-2-mediated ErbB activity
(Bahrami et al., 2010). In contrast, endogenous GLP-2 action does not ameliorate the severity
and extent of large bowel injury in experimental colitis, consistent with low GLP-2R density
in the colon (Lee et al., 2012).
GLP-2R KO mice do not exhibit disrupted epithelial barrier function under normal
conditions. However, GLP-2R KO mice are more susceptible to enteritis-induced bacterial
translocation and morbidity (Lee et al., 2012). One potential explanation for this finding is
that endogenous GLP-2 is required for the maintenance of barrier function under disease
states, such as small intestinal injury triggered by chemotherapeutic irinotecan or the non-
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steroidal inflammatory drug indomethacin. Additionally, under healthy conditions, GLP-2R
deletion alters the intestinal microbiota composition and perturbs Paneth cell function by
reducing bactericidal activity and antimicrobial gene expression, both of which may
contribute to the higher sensitivity to small bowel injury observed in GLP-2R KO mice (Lee
et al., 2012).
1.3.2 Actions of exogenous GLP-2
Pharmacologic effects of GLP-2, mostly determined using the long-lasting analogue
Gly2GLP-2, have been extensively studied. Exogenous GLP-2 has been determined to exert a
plethora of effects on GI growth and function, as well as exhibiting cytoprotective effects to
ameliorate intestinal injury.
1.3.2.1 Epithelial barrier function
An abundance of evidence shows that GLP-2 improves intestinal barrier function.
The first study was elegantly conducted by Benjamin et al. to show that GLP-2-mediated
enhancement of the epithelial barrier occurs through both paracellular and transcellular
pathways (Benjamin et al., 2000). Chronic 10-day treatment of mice with either native GLP-
2 or Gly2GLP-2 reduced paracellular permeability, as measured by ion conductance and
passage of a radiolabelled inert probe, 51
Cr-EDTA. Through the transcellular pathway, flux
of horseradish peroxidase (HRP) was also reduced. Importantly, acute administration of
Gly2GLP-2 is also sufficient to reduce intestinal permeability, as measured by all three
parameters mentioned above, within 4 or 48 hr. However, the decrease in permeability
cannot be merely explained by increased paracellular transit time since enterocyte length was
15
not increased at 48 hr. This suggests involvement of tight junctions in GLP-2-reduced
paracellular permeability.
Under conditions of barrier dysfunction, as demonstrated by models of acute
necrotizing pancreatitis, food allergy and stress, GLP-2 treatment also reduces intestinal
permeability, and this effect is accompanied by amelioration of impaired host defense
(Cameron and Perdue, 2005; Cameron et al., 2003; Kouris et al., 2001). Restoration of barrier
function in murine models of enteritis is also suggested by reduced bacterial translocation
and increased survival rate with GLP-2 administration (Boushey et al., 1999; Boushey et al.,
2001). Recently, GLP-2 has also been shown to reduce gut permeability in the presence of
low-grade inflammation, as reported in obese leptin-deficient mice, which is suggested to be
associated with changes within tight junction complexes (Cani et al., 2009). Importantly, a
downstream mediator responsible for the enhanced barrier effects of GLP-2 has not been
identified to date.
1.3.2.2 Mucosal growth
The most profound and well-characterized aspect of GLP-2 actions is induction of
mucosal growth. GLP-2 increases small and large intestinal weight through stimulation of
epithelial cell proliferation and inhibition of apoptosis, resulting in increased crypt depth as
well as villus and microvillus height, thereby enlarging absorptive surface area (Drucker et
al., 1996; Drucker et al., 1997a; Tsai et al., 1997a; Tsai et al., 1997b; Benjamin et al., 2000).
Trophic effects of GLP-2 are most evident in the jejunum and least pronounced in the colon,
consistent with GLP-2R distribution (Drucker et al., 1999; Dube et al., 2006; Munroe et al.,
1999; Yusta et al., 2000). GLP-2 growth effects are reported across various mammalian
16
species, including humans (Jeppesen et al., 2005), pigs (Burrin et al., 2000; Pereira-Fantini et
al., 2008), rats (Drucker et al., 1997a) and mice (Benjamin et al., 2000; Drucker et al., 1996).
Furthermore, GLP-2-induced mucosal growth occurs in association with cytoprotective
effects accompanied by attenuation of the extent of intestinal injury in various disease
models including, but not limited to, enteritis (Boushey et al., 1999; Boushey et al., 2001)
and colitis (Drucker et al., 1999).
The key players mediating the growth effects of GLP-2 identified to date are IGF-1
and ErbB ligands in the small intestine, and keratinocyte growth factor (KGF) in the colon
(Orskov et al., 2005; Dube et al., 2006; Yusta et al., 2009). GLP-2 increases IGF-1 mRNA
levels in intestinal SEMFs (Leen et al., 2011), and IGF-1 and, to a lesser extent, IGF-2 are
required for the trophic effects of GLP-2 using global murine gene deletion models (Dube et
al., 2006; Dube et al., 2008). Furthermore, conditional deletion of the IGF-1R specifically in
the intestinal epithelium of adult mice results in reduced proliferative and growth effects to
GLP-2. Similarly, the ErbB ligands, epiregulin and neuregulin, are upregulated in the murine
small intestine in response to GLP-2 stimulation (Bahrami et al., 2010), and ErbB receptors
are required for the intestinal proliferative response to GLP-2 (Yusta et al., 2009). It has been
suggested that the IGF-1/IGF-1R and ErbB ligand/ErbB signaling pathways may interact.
GLP-2 does not affect ErbB ligand mRNA transcript levels in SEMFs (Leen et al., 2011),
and epidermal growth factor (EGF), but not IGF-1, can restore growth responses in refed
GLP-2R KO mice (Bahrami et al., 2010). Taken together, these findings suggest that IGF-1
resides upstream of ErbB ligand/ErbB activation. Importantly, transactivation between the
IGF-1R and ErbB is also well-established, demonstrating cross-talk between these two
17
pathways (Ahmad et al., 2004; Gschwind et al., 2001; Jin and Esteva, 2008; Roudabush et
al., 2000).
1.3.2.3 Other intestinal functions
In addition to barrier and growth effects, GLP-2 exerts complementary actions that
are beneficial to GI function. GLP-2 rapidly enhances hexose absorption through stimulation
of GLUT-2 activity and upregulation of SGLT-1 (Cheeseman, 1997; Ramsanahie et al.,
2003). Absorption of triglycerides and amino acids is also elevated with chronic GLP-2
treatment (Brubaker et al., 1997b; Kato et al., 1999; Scott et al., 1998). Furthermore, chronic
administration of GLP-2 enhances disaccharide digestion by increasing activity and gene
expression of digestive enzymes (Brubaker et al., 1997b; Kitchen et al., 2000; Petersen et al.,
2002). Moreover, enhanced digestion and absorption can be coupled with GLP-2 induced
stimulation of small intestinal blood flow through endothelial nitric oxide synthase (Guan et
al., 2006). GLP-2 also inhibits antral motility (Nagell et al., 2004; Wojdemann et al., 1998),
which may contribute to the ileal brake effect whereby lengthened transit time through the GI
tract increases nutrient exposure to facilitate absorption. Lastly, the intestinal lumen is under
constant exposure to bacterial antigens, and GLP-2 has been demonstrated to have anti-
inflammatory effects mediated through vasoactive intestinal peptide (VIP) (Sigalet et al.,
2007).
1.4 Insulin-like growth factors
The IGF system consists of the two ligands, IGF-1 and IGF-2, which share 70%
amino acid sequence homology, the receptors, IGF-1R and IGF-2R (also known as the
18
mannose-6-phosphate R), and seven IGF binding proteins, IGFBP-1 to -7. The IGFs are
peptide hormones known to promote development and their actions overlap with many of
GLP-2 functions in the GI tract, including promoting growth and barrier function (Benjamin
et al., 2000; Dube et al., 2006; Lorenzo-Zuniga et al., 2006; Peterson et al., 1996; Rowland et
al., 2011). The Brubaker laboratory has shown that GLP-2 signals upstream of the IGF and
IE-IGF-1R system in the gut to promote epithelial proliferation (Dube et al., 2006; Rowland
et al., 2011), which raises the possibility that this pathway may be involved in transducing
other effects of GLP-2 as well.
1.4.1 IGF peptides
The insulin-like growth factors were discovered as mediators of growth hormone-
induced skeletal growth (Daughaday et al., 1987). The name comes from recognition that
these peptides are structurally homologous to insulin (Rinderknecht and Humbel, 1978; Chan
et al., 1992). IGF-1 and IGF-2 are insulin-like in that their A and B domains are connected by
disulfide bonds homologous to those of insulin. However, unlike proinsulin, IGFs do not
undergo PC1/3 or PC2-mediated proteolytic cleavage, and remain connected by the C
domain. In addition, the IGFs contain an extra D domain not found in proinsulin (Chan et al.,
1992; Rinderknecht and Humbel, 1978).
IGF-1 is expressed and secreted by virtually all tissues, and its sites of secretion often
reflect its actions (Laron, 2001). The liver is the major source of circulating IGFs; whereas
IGF-1 is under the control of the hypothalamic-pituitary-liver axis, which regulates post-natal
and pubertal growth, IGF-2 is released from the liver in a constitutive manner (Efstratiadis,
1998). Fetal growth in early gestation is predominantly regulated by IGF-2 with IGF-1 levels
19
on the rise close to full gestation (Efstratiadis, 1998). IGFs are also released by the GI tract,
in which autocrine and paracrine mechanisms play a vital role (Liu et al., 2000; Sjogren et
al., 1999). In the intestine, IGF-1 is expressed primarily by two cell types, the SEMFs and the
smooth myocytes (MacDonald, 1999; Ohneda et al., 1997; Yakar et al., 2005).
1.4.2 Metabolism and clearance
Upon secretion, IGFs can be bound to the IGFBPs to modulate their plasma half-lives
and potency (Firth and Baxter, 2002). IGFBPs serve as a reservoir for IGFs through
sequestration and regulated release of IGFs, and can thereby potentiate as well as inhibit IGF
activity (Pollak et al., 2004). In particular, IGFBP-3, -4 and -5 predominate in the rodent and
human postnatal intestine, and IGFBP-3, in particular, is the most prominent and has the
highest binding capacity in humans (Lund, 1998). In addition, the IGF-IGFBP dimer can
bind to an 85-kDa acid-labile subunit to form a large 150kDa multicomplex, which can
further prolong IGF half-life (Boisclair et al., 2001). IGFBP activity is regulated by protease
digestion (Pollak et al., 2004). Renal clearance of the IGFs is rapid when not bound to
IGFBPs and results in a half-life of 2 min in the SI (Xian et al., 1995). Degradation of the
IGF peptides requires IGF-1R binding (Pollak et al., 2004), and subsequent digestion by
proteolytic insulin-degrading enzyme (Misbin and Almira, 1989).
1.4.3 The IGF-1R
Similar to the IGF peptides, the IGF-1R is also expressed by virtually all tissues, and has
been localized to the GI mucosa and muscularis in particular (Howarth, 2003; Laburthe et al.,
1988). In the mucosal layer, the IGF-1R is expressed in crypt and villus enterocytes with
20
higher levels in the crypt (Laburthe et al., 1988; Ney et al., 1999). The receptor is localized to
both the apical and basolateral domains of the cell membrane but exhibit higher density on
the basolateral side of the enterocyte. Both IGF-1 and -2 are ligands for the IGF-1R, which is
similar in structure to the insulin receptor (IR; Figure 1.3). The IGF-1R and IR are both
integral receptors belonging to the growth factor tyrosine kinase receptor family (Inagaki et
al., 2007; Ward et al., 2001). These receptors are composed of two extracellular α subunits
involved in ligand binding and two transmembrane β subunit with intrinsic tyrosine kinase
activity. Encoded on chromosome 15, exons 1-10 of IGF-1R make up the α-subunit wherein
exon 3 is essential for binding capacity, and exons 12-21 encoding the β-subunit (Inagaki et
al., 2007; Ward et al., 2001). The extracellular ligand binding α subunit of the IGF-1R
contains two leucine-rich repeat domains L1 and L2 intervened by a cysteine-rich region,
whereas the transmembrane signaling β subunit is tyrosine-rich and contains a domain with
tyrosine kinase activity. Since IGF-1R and IR share high structural similarity, hybrid
receptors containing the IGF-1R αβ dimer bound to the IR αβ dimer can be found in cells
expressing both receptors (Inagaki et al., 2007; Ward et al., 2001).
21
Figure 1.3 Structure and intracellular signaling of the IGF-1R.
The IGF-1R is heterotetrameric comprised of two extracellular α chains and two
transmembrane β chains. Each α chain contains two leucine-rich domains, L1 and L2, as well
as an intervening cysteine-rich domain, which contains exon 3 and is important for IGF
binding. The β chain contains a tyrosine kinase domain and undergoes autocatalytic
phosphorylation, which leads to activation of IRS. Downstream recruitment of p85 subunit of
PI3K leads to activation of the AKT pathway, and recruitment of Grb2 results in activation of
22
the Erk pathway. IGF-1R intracellular signal transduction promotes cell growth,
differentiation and survival.
23
Upon ligand binding, the IGF-1R undergoes autocatalytic phosphorylation at Y1131,
1135, 1136 that induces further phosphorylation at Y
943, 950 , 1316 (Kato et al., 1994; Li et al.,
1994). Tyrosine phosphorylation of the IGF-1R opens up docking sites for recruitment
signaling molecules containing phosphotyrosine binding domains, such as insulin receptor
substrate (IRS)-1 to -4 and the Src-homology adaptor protein (Shc) (Craparo et al., 1995a;
Dey et al., 1996; Tartare-Deckert et al., 1995; Xu et al., 1999). Subsequent phosphorylation
of IRS protein provides additional docking sites for downstream molecules, such as PI3K and
Grb2 (Craparo et al., 1995b; Myers, Jr. et al., 1993; Sun et al., 1991).
Downstream of IRS signaling, the PI3K pathway activation starts with the p85
subunit of PI3K binding to IRS, which activates its p110 subunit (Shepherd et al., 1998).
Subsequently, phosphatidylinositol-(4,5)-bisphosphate (PIP2) is converted to
phosphatidylinositol-(3,4,5)-triphosphate (PIP3). This then leads to activation of the
pleiotrophic growth and survival enzyme, Akt (Alessi et al., 1996).
Another pathway activated by IGF-1R signaling is the Erk pathway, also known for
effects of proliferation and survival. The MAPK signaling cascade is initiated by recruitment
of the Grb2 adapter protein and the GTP-exchange factor SOS by IRS or by Shc association
with the IGF-1R (Ravichandran, 2001). Downstream signaling involves sequential
phosphorylation and activation of protein kinases – the G protein Ras, Raf, Mek1/2, then
Erk1/2. The Erk1/2 enzymes are known to regulate translation, mitosis and apoptosis (Yoon
and Seger, 2006). Collectively, IGF-1R signaling mediates cellular pathways involved in
growth, differentiation and survival.
24
1.5 Epithelial barrier function
The intestinal mucosal barrier maintains an intricate balance between absorption of
nutrients and prevention of pathogens from entering the circulation. It is comprised of the
outer and inner mucus layers, the IE monolayer, as well as the inner subepithelial innate and
adaptive immune systems (Salim and Soderholm, 2011; Turner, 2009; Rescigno, 2011). The
IE cells form a continuous and polarized physical barrier, linked together by a series of
dynamic apical junction complexes that serve as the integral component of the mucosal
barrier and tend to be the rate-limiting factor in passage via the paracellular pathway. Low
molecular weight molecules cross this barrier through the paracellular route, whereas larger
macromolecules may transit by transcellular transport (Watson et al., 2001). The junctional
complexes consist of desmosomes, gap junctions, adherens junctions, and the most
luminally-situated, tight junctions that serve as the first barrier in this dynamic network
(Figure 1.4).
25
A
B
Figure 1.4 Intestinal epithelial apical junction complexes.
Molecules can pass through the epithelial monolayer through the transcellular pathway that is
comprised of tight junctions, adherens junctions, gap junctions and desmosomes. The plasma
26
membranes of adjacent cells are fused just below the base of the microvilli at the tight
junction. Junctional proteins consist of claudins, occluden and zona occludens (ZOs). ZOs
are connected to intracellular F-actin.
27
The anastomosing network of the tight junctional complex is comprise of
transmembrane proteins (claudins, occludin, and junctional adhesion molecules; JAMs),
peripheral scaffolding proteins (zona occludens; ZOs), and intracellular regulatory molecules
(kinases and actin; Figure 1.5) (Turner, 2009; Hossain and Hirata, 2008; Bruewer et al.,
2006). The integral proteins, claudins and occludin, can homodimerize between adjacent
cells to form seals or pores in the paracellular space, as well as directly interact with
intracellular scaffolds. The family of claudins consists of 24 members, which are tetraspan
membrane proteins of ~20 kDa in size, and that primarily determine paracellular permeability
based on charge and size selectivity (Furuse and Tsukita, 2006; Lal-Nag and Morin, 2009;
Tsukita et al., 2001). Particular claudins can also serve as tightening or pore-forming
junctional components. Transmembrane occludin is also tetraspan, but larger in size (65 kDa
in its unphosphorylated form), and is known to be a sealing protein (Cummins, 2012). The
family of JAMs belongs to the immunoglobulin superfamily and is involved in epithelial
barrier function as well as cell-to-cell adhesion of endothelial cells (Martin-Padura et al.,
1998). A newly recognized member of the tight junction complex, tricellulin, is involved in
forming intercellular links between three neighboring cells (Ikenouchi et al., 2005; Krug et
al., 2009). In contrast to bicellular tight junctions, tricellulins are too rare to significantly
contribute to ion permeability, although they can permit the passage of macromolecules
(Krug et al., 2009). Finally, the peripheral membrane proteins, ZO-1, -2 and-3, play a vital
role in tight junctional complex formation by connecting the strand-forming tight junction
proteins with the cytoskeletal actin microfilaments. These ZOs are regulated by various
intracellular signaling pathway effectors, such as myosin light chain kinase, and thereby
28
modulate the assembly, maintenance, and barrier function of the tight junction complex
(Schulzke et al., 2005).
GLP-2 has been linked to modulation of tight junctional proteins in the maintenance
of the epithelial barrier. In the ob/ob mouse model, administration of a prebiotic increases
junctional proteins, ZO-1 and occludin, at the mRNA level, and these effects are GLP-2-
dependent as shown by antagonist studies (Cani et al., 2009). In addition, studies using a
Caco-2 cell model demonstrate that GLP-2 reduces trans-epithelial conductance in
association with up-regulation of ZO-1 and occludin (Moran et al., 2012), although it remains
unclear as to whether these cells actually express the GLP-2R (Yusta et al., 2000).
Nonetheless, knowledge of GLP-2 signalling in enhancing barrier function remains very
limited.
1.6 Hypothesis and specific aims
The growth factors GLP-2 and IGF-1 have both been implicated in reducing
transepithelial permeability (Huang et al., 1993; Alexandrides et al., 1998; Lorenzo-Zuniga et
al., 2006; Benjamin et al., 2000). However, whether these two growth factors interact or
participate in the same pathway remain uncertain. Based on GLP-2R distribution, it is likely
that GLP-2-induced reductions in permeability require a downstream mediator and, thus, the
hypothesis of the current thesis is that the GLP-2-mediated effects on barrier function require
the IE-IGF-1R-dependent pathway. To interrogate this hypothesis, the IE-IGF-1R was
targeted using an inducible knockout mouse model, and was investigated under models of
health and enteritis. The specific aims of this thesis were to determine the role of the IE-IGF-
29
1R in GLP-2-induced barrier effects: (i) in vivo by measuring GI permeability, (ii) ex vivo by
measuring jejunal resistance, and (iii) by examining subcellular changes in jejunal tight
junction proteins at the levels of expression and localization.
30
2 MATERIALS AND METHODS
2.1 Animals
All animals were bred and housed in a 12-hour light/dark cycle animal facility at the
University of Toronto. All animal studies were approved by the University of Toronto
Animal Care Committee. IE-IGF-1R KO mice were generated by crossing villin-CreERT2+/0
(from Dr. S. Robine) and Igf1rflox/flox
mice (from Dr. M. Holzenberger via Dr. R.N. Kulkarni)
(Desbois-Mouthon et al., 2006; el et al., 2004; Kappeler et al., 2008; Rowland et al., 2011),
both on a C57BL/6 background. The villin-CreERT2+/0
; Igf1rflox/+
offspring were then
backcrossed to Igf1rflox/flox
mice to generate the villin-CreERT2+/0
; Igf1rflox/flox
animals, named
the IE-IGF-1R KO mice. Mice were genotyped as previously described (Leneuve et al.,
2001; Rowland et al., 2011). Briefly, control mice were identified by detection of the floxed
allele, whereas IE-IGF-1R KO mice were identified by detection of the additional Cre allele.
Oligonucleotides 5’-ATCTTGGAGTGGTTGGGTCTGTTT-3’ and 5’-
ATGAATGCTGGTGAGGGTTGTCTT-3’ amplified a 327-bp fragment of the floxed allele,
and the primers 5’-CCTGGAAAATGCTTCTGTCCG-3’ and 5’-
CAGGGTGTTATAAGCAATCCCC-3’ amplified a 390-bp fragment from the Cre coding
region (Figure 2.1). Age- and sex-matched littermate IE-IGF-1R KO and Igf1rflox/flox
control
mice were used in all experiments.
31
Figure 2.1 Generation of the IE-IGF-1R KO mouse using the Cre/lox system
Villin-CreERT2+/0
and igf1rflox/flox
mice were crossed to generate the villin-CreERT2+/0
;
igf1rflox/flox
(IE-IGF-1R KO) animals. Exon 3, required for ligand binding, was deleted by
tamoxifen induction. Fragments of the floxed allele and Cre coding region were detected by
PCR at 327 and 390 bp, respectively.
32
IE-IGF-1R exon 3 cleavage was induced by nuclear translocation of Cre recombinase
in mice aged 8-13 weeks, by daily intraperitoneal injection of tamoxifen (100 µl of 10
mg/ml; reconstituted in ethanol at 100 mg/ml and diluted in sunflower oil; MP Biomedicals,
Solon, OH) for 5 days (el Marjou et al., 2004; Rowland et al., 2011). Tamoxifen was also
administered to Igf1rflox/flox
mice to control for tamoxifen side-effects. After tamoxifen
induction, three experimental protocols were followed for animal models of health,
irinotecan-induced enteritis and streptocozotocin-induced diabetes. Since the particular
streptozotocin protocol used only induced glucose intolerance, it is therefore described in
detail in the Appendix instead. For the healthy animal model, data from female and male
mice were combined. Animals were injected subcutaneously with GLP-2 (0.1 µg/g
h(Gly2)GLP-2; American Peptide Company; Sunnyvale, CA) or vehicle (phosphate-buffered
saline [PBS]) daily for 10 days, with the final booster injection 3 hr before permeability
assessment. For the enteritis model, only males were included because of higher sensitivity to
irinotecan treatment, as determined in preliminary studies (Figure 2.2). Animals received 10
days of GLP-2 or vehicle treatment, followed by intraperitoneal injection of irinotecan (0.15
µg/g body weight; reconstituted in ethanol at 0.1 g/ml and diluted in PBS; Sigma-Aldrich,
Oakville, ON) for two days. After 3 days of recovery, in vivo and ex vivo permeability
assays were performed on day 4 and day 6, respectively.
33
Figure 2.2 Male mice appeared more sensitive to irinotecan than female mice.
Intestinal FD4 permeability was measured to determine the effect of irinotecan on barrier
function in male and female mice (n=3-4).
34
Mice were anesthetized with isofluorane prior to sacrifice by cervical dislocation and
the small intestine was removed and cleaned of luminal content. A 2-cm segment from the
mid-jejunum was fixed in 10% neutral-buffered formalin overnight before paraffin
embedding and sectioning (at Toronto General Hospital pathology lab). A 2-mm section
proximal to the mid-piece was cut in half for electron microscopy following fixation in 2.5%
glutaraldehyde overnight. An additional 2-cm segment was collected distal to the mid-piece
and immediately frozen on dry ice for immunoblotting.
2.2 Intestinal permeability
GI permeability was measured in vivo using the relatively impermeant 4 kDa
macromolecule, fluorescein isothiocyanate dextran (FD4; Sigma-Aldrich). Mice were fasted
overnight and orally gavaged with FD4 (0.5 mg/g; 50 mg/ml PBS). After 1.5 hr, 120 µl of
tail vein blood was collected and centrifuged at 13000g for 5 min. In a 96-well plate, 50 µl of
plasma was diluted with an equal volume of PBS, and standard curves were generated by
diluting FD4 in an equal volume of non-treated plasma. FD4 fluorescence was measured at
an excitation wavelength of 485 nm and emission wavelength of 535nm.
Transmural resistance of two contiguous segments of mid-jejunum was measured ex
vivo in Ussing chambers. In brief, 5-mm segments were opened along the mesenteric border
and mounted into Ussing chambers (Physiologic Instruments, San Diego, CA). Tissues were
incubated in modified Meyler solution (128 mM NaCl, 4.7 mM KCl, 1 mM MgCl2, 0.3 mM
Na2HPO4, 0.4 mM NaH2PO4, 20 mM NaHCO3, 10 mM HEPES, 1.3 mM CaCl2, pH 7.3) at
37°C with continuous oxygenation. After equilibration (~20min on average), tissues were
35
clamped with intermittent current pulses of 0.001 mA, and the corresponding changes in
voltage were continuously recorded. Resistance was calculated according to Ohm’s law using
values of applied current and the resultant potential difference (R=ΔV/ΔI). Integrity of the
tissue was tested using forskolin (10 µM; Sigma-Aldrich) plus 3-isobutyl-1-methylxanthine
(200 µM; Sigma-Aldrich) to induce cystic fibrosis transmembrane conductance regulator-
mediated chloride secretion; tissues with no response were considered damaged and were
excluded from analyses.
2.3 Microscopy
Crypt-villus height was measured on hematoxylin and eosin (H&E)-stained slides. An
average length of 38 well-oriented villi with intact crypts from 3-4 cross sections per mouse
was quantified. For electron microscopy, tissues underwent routine processing at Division of
Pathology, Hospital for Sick Children, and photomicrographs of well-oriented epithelial cells
were taken and evaluated with Dr. Cameron Ackerley.
For immunostaining, 4-µm jejunal cross sections were prepared from formalin-fixed,
paraffin-embedded tissues. Antigen retrieval for ZO-1 and occludin immunostaining was
protease-induced, with pronase E (1 mg/ml in 0.05M Tris buffer, pH 7.6; Sigma-Aldrich)
incubation for 15 min at 37°C, whereas for claudins, it was heat-induced by microwaving in
sodium citrate buffer (10 mM, pH 6; Sigma-Aldrich) for 20 min. Sections were then washed
with two changes of PBS, blocked with 10% normal serum for 1.5 hr at room temperature,
and incubated overnight at 4°C with primary antibodies; rabbit anti-ZO-1 (1:100; Invitrogen,
Camarillo, CA), -occludin (1:125; Invitrogen), -claudin-3 (1:40; Abcam, Cambridge, MA), -
claudin-7 (1:100; Invitrogen), -claudin-15 (1:20; Invitrogen), and goat anti-sucrase (1:150;
36
Santa Cruz Biotechnology, Santa Cruz, CA). Following probing with primary antibody,
slides were washed in Tris-buffered saline/0.1% Tween 20 (v/v; TBST) three times at 10 min
intervals. Secondary antibody incubation was performed at room temperature for 1 hr. ZO-1
and occludin were detected using Alexa Fluor 555 goat anti-rabbit IgG (1:500; Invitrogen),
the claudins by CY3 donkey anti-rabbit (1:200; Jackson ImmunoResearch Laboratories,
West Grove, PA), and sucrase by FITC donkey anti-goat IgG (1:200; Jackson
ImmunoResearch Laboratories). An AxioPlan deconvolution microscope (Carl Zeiss,
Canada, Don Mills, ON) was used to acquire all images, and exposure levels were fixed
constant for every slide, which contained tissues from 4 different animals, consisting of one
from each treatment group subject to comparison.
2.4 Immunoblotting
Jejunal mucosa was scraped following freeze-fracturing and homogenized in RIPA
lysis buffer (50 mM β-glycerol phosphate, 10 mM Hepes, 1% Triton X-100, 70 mM sodium
chloride, 2 mM EGTA, 1mM sodium orthovanadate, 1 mM sodium fluoride and 1 complete
mini EDTA-free protease inhibitor tablet (Roche Diagnostics Corp., Indianapolis, IN).
Protein concentration was quantified by Bradford assay (Bio-Rad, Hercules, CA), and 100 µg
of protein per sample was loaded onto a 7% (for ZO-1 and occludin) or 15% (for claudins)
polyacrylamide gel. Protein samples were transferred at 4°C for 1.5h at 110 V or overnight at
30 V onto an Immun-Blot PVDF membrane (Bio-Rad). The membrane was probed with
rabbit anti-ZO-1 (1:200), -occludin (1:200), -claudin-3 (1:2000), -claudin-7 (1:4000), -
claudin-15 (1:1000), or –β actin, loading control (1:4000; Sigma-Aldrich). Subsequently, the
immunoblot was washed 5 times with TBST at 5 min intervals, and was probed with
37
horseradish peroxidase-linked goat anti-rabbit IgG (1:2000; Cell Signaling Technology,
Beverly, MA). After another 5 x 5 min washes, bands were detected with ECL Western
blotting detection reagent (Amersham GE Healthcare, Baie d’Urfe, QC). The membrane was
visualized using Kodak imager 4000pro (Carestream, Rochester, NY).
2.5 Statistical analyses
All data are expressed as mean ± standard error. Results were analyzed by two-way
analysis of variance (ANOVA), followed by Student’s t test. P values of <0.05 were
considered statistically significant.
38
3 RESULTS
3.1 Validation of the IE-IGF-1R KO mouse model
To investigate the GLP-2 signalling pathway in enhancing barrier function, the inducible IE-
specific IGF-1R KO mouse model was utilized. IGF-1Rfl/fl
control and IE-IGF-1R KO
littermates were identified by genotyping (Figure 3.1A). PCR analysis demonstrated the
presence of the floxed allele in both control and KO animals, whereas the Cre allele was
present only in the KO animals. After treating both control and KO animals with tamoxifen,
functional validation of gene deletion was performed by assessment of GLP-2-induced
jejunal crypt-villus growth (Figure 3.1B), which is known to require the IE-IGF-1R
(Rowland et al., 2011). In response to GLP-2 treatment, control mice demonstrated an
increase in crypt-to-villus height, by 27.6 ± 2.9 % (p<0.01). In contrast, crypt-to-villus height
did not differ between vehicle- and GLP-2-treated KO mice.
39
Figure 3.1 Identification and validation of the IE-IGF-1R KO mouse model.
(A) Genotype-specific primers were used to amplify the floxed and Cre alleles, generating
bands of 327 and 390 bp, respectively. (L = ladder; - = empty) (B) Jejunal crypt-to-villus
height was measured in control and KO mice following treatment for 10 d with vehicle or
GLP-2 (n=4-8, **p<0.01).
40
3.2 The IE-IGF-1R was essential for GLP-2-enhanced barrier function
To determine the in vivo effects of chronic GLP-2 treatment on permeability along
the GI tract, fluorescent macromolecule FD4 levels were measured in the blood following
oral gavage. In control mice, plasma FD4 levels were reduced with GLP-2 treatment, by 61.9
± 7.82 % (p<0.05; Figure 3.2A). FD4 levels were not changed in vehicle-treated KO mice.
However, this GLP-2 response of reduced FD4 permeability was abolished in the absence of
the IE-IGF-1R, demonstrating that GLP-2-lowered GI permeability is dependent upon the IE-
IGF-1R.
Since the GLP-2R is expressed at the highest levels in the jejunal segment of the
small bowel (Munroe et al., 1999; Yusta et al., 2000), ex vivo transmural tissue resistance
was quantified in jejunal tissue with Ussing chamber measurements. GLP-2 treatment
increased resistance, by 44.1 ± 6.0 % (p<0.001), in control mice, as compared to 22.3 ± 8.2
% (p=ns) in KO animals (Figure 3.2B). Basal levels of resistance were not statistically
different between vehicle-treated control and KO mice (p=0.27). The results confirm that, at
the jejunal level, GLP-2 enhanced barrier function and required the IE-IGF-1R.
3.3 No change in tight junctional complex ultrastructure was detected by
electron microscopy
To examine the ultrastructure of the tight junctional complex, transmission electron
photomicrographs were taken. No changes were detected between the four treatment groups
(Figure 3.3).
41
Figure 3.2 GLP-2-enhanced barrier function was reduced in KO mice.
(A) Circulating levels of FD4, a relatively impermeant fluorescent marker, were measured
following oral gavage (n=11-14, *p<0.05). (B) Jejunal tissue resistance was quantified in
42
Ussing chambers (n=6-9, ***p<0.001). Vehicle and GLP-2 treatments are presented by white
and black bars, respectively.
43
Figure 3.3 Ultrastructure of tight junctional proteins was unchanged.
Representative electron micrographs were taken from mid-jejunum of (A) vehicle-, and (B)
GLP-2-treated control mice, (C) vehicle-, and (D) GLP-2-treated KO mice (n=3).
44
3.4 GLP-2 upregulated claudin-3 and -7 expression via the IE-IGF-1R
Selected tight junctional proteins that regulate jejunal paracellular permeability were
analyzed to determine possible mechanisms underlying the observed increase in barrier
function stimulated by GLP-2. Claudin-3 was upregulated by GLP-2 treatment, by 4.6 ± 1.7
fold (p<0.05), in control animals (Figure 3.4A). However, this GLP-2-induced expression of
claudin-3 was abrogated in KO animals. Similarly, claudin-7 was upregulated in response to
GLP-2 by 0.5 ± 0.2 fold (p<0.01) in control, but not KO animals (Figure 3.4B). There were
no changes in basal expression levels of either claudins between control and KO mice. These
findings were consistent with the increased immunofluorescence intensity observed in
claudin-3 and -7 immuno-stainings in control but not KO mice (Figure 3.5). In contrast,
pore-forming claudin-15 expression appeared unchanged in all groups of animals (Figure
3.4C & 3.5). In addition, no changes in subcellular localization were detected for any of the
claudins examined; claudin-3, -7 and -15 were localized to the cellular membrane region, as
opposed to intracellular domains. Taken together, these findings indicate that GLP-2
upregulated claudin-3 and -7 expression in an IE-IGF-1R-dependent manner.
45
Figure 3.4 Protein expression levels of tight junctional claudins.
(A) Claudin (cl)-3 (n=6-9), (B) cl-7 (n=8-9) and (C) Cl-15 (n=5-7; additional n values
pending) expression levels were quantified by immunoblotting (*p<0.05, **p<0.01). Vehicle
and GLP-2 treatments are presented by white and black bars, respectively.
46
47
Figure 3.5 Immunofluorescence of claudin-3, -7 and -15.
Claudins were immunostained in red, the brushborder marker sucrase in green, and nuclei in
blue (n=4); both longitudinal views of the villi and tight junctional webbing patterns are
shown for each claudin. Pictures subjected to comparison were captured under the same
exposure settings. Negative control staining was done without primary but with the same
secondary antibody used for cl-3, -7 and -15 staining.
48
3.5 Deletion of the IE-IGF-1R altered occludin localization
Occludin total protein expression levels were not different between the four groups of
animals (Figure 3.6A). Immunostaining of the junctional sealing protein, occludin, revealed
that, under basal conditions, occludin is localized predominantly along the apical border as
seen in vehicle-treated control mice (Figure 3.7). In response to GLP-2 treatment, control
animals appeared to have more prominent apical occludin localization. In contrast, vehicle-
treated KO animals exhibited more intracellularly-situated occludin, and GLP-2 treatment
did not induce the apical localization of occludin in KO animals. Thus, the results suggest
that the IE-IGF-1R is required for occludin localization, suggestive of integration into the
tight junction complex.
3.6 Scaffolding ZO-1 was unchanged with IE-IGF-1R deletion and/or
GLP-2 treatment
ZO-1 is an adaptor protein that connects transmembrane junctional proteins to
cytoskeletal actin (Salim and Soderholm, 2011). Immunoblotting and immunostaining results
both showed that ZO-1 expression and localization did not change with GLP-2 stimulation
and were unaltered in the IE-IGF-1R KO animals (Figure 3.6B & 3.7). These findings
suggest that changes in ZO-1 may not be involved in GLP-2-enhanced barrier function.
49
Figure 3.6 Protein expression levels of occludin and ZO-1.
(A) ZO-1 (n=6-8) and (B) occludin (n=10-11) protein levels were quantified by
immunoblotting. White bars and black bars represent vehicle- and GLP-2-treated mice,
respectively.
50
Figure 3.7 Subcellular localization of occludin and ZO-1.
Occludin and ZO-1 were immunostained in red and nuclei in blue (n=4). Pictures subjected
to comparison were captured under the same exposure settings. Negative control staining was
conducted without primary and with the respective secondary antibody.
51
3.7 The IE-IGF-1R was required for GLP-2-normalized barrier function
in irinotecan-induced enteritis
To determine the barrier effects of GLP-2 under conditions of mucosal inflammation,
chemotherapeutic irinotecan was used to chemically-induce enteritis. Irinotecan treatment
resulted in pathologic patchy morphology, with patchy areas of inflammation and villus
damage (Figure 3.8A-D). Residual intact villi and crypt were quantified, and irinotecan-
treated control, but not KO, mice remained responsive to the growth effects of GLP-2,
consistent with findings in healthy animals (Figure 3.8E). It was expected that irinotecan
would disrupt barrier function in control mice (Lee et al., 2012; Boushey et al., 2001), and
that GLP-2 would restore barrier function, based on findings in healthy mice (Figure 3.2).
However, unexpectedly after irinotecan induction, GLP-2-treated control animals had
increased GI permeability to FD4 (Figure 3.9A) and reduced jejunal resistance (Figure
3.9B) as compared to vehicle-treated control animals with irinotecan-induced enteritis.
Notwithstanding, the effects of GLP-2 on GI permeability and jejunal resistance were
reduced or absent in the KO mice, respectively. These findings suggest that GLP-2
normalized intestinal permeability in this model of altered barrier function, and this effect
required the IE-IGF-1R.
52
53
E
Figure 3.8 GLP-2-restored crypt-villus growth in irinotecan-induced enteritis required
the IE-IGF-1R.
Representative examples of H&E staining of jejunal tissue from irinotecan-treated mice in
(A) vehicle-treated control, (B) GLP-2-treated control, (C) vehicle-treated KO, (D) GLP-2-
treated KO animals are shown. (E) Height of the crypt-villus axis was measured in
irinotecan-treated mice (n=7-10). Dashed line represents vehicle-treated healthy animals,
from Figure 3.1B. White bars and black bars represent vehicle- and GLP-2-treated mice,
respectively (*p<0.05; **p<0.01).
54
A B
Figure 3.9 GLP-2-normalized barrier function required the IE-IGF-1R in irinotecan-
induced enteritis
Barrier function was measured by (A) plasma FD4 levels (n=8-11) and (B) jejunal transmural
resistance (n=5-9). Dashed lines represent values of vehicle-treated healthy animals, from
previous figures. White bars and black bars represent vehicle- and GLP-2-treated mice,
respectively (*p<0.05).
55
3.8 GLP-2-normalized barrier function may require mechanisms
independent of the tight junctions
Tight junction proteins were examined following induction of enteritis with irinotecan
by both western blot and immunostaining of jejunal tissues. In contrast to healthy mice, GLP-
2 did not up-regulate protein expression levels of claudin-3 and -7 in mice with enteritis.
Interestingly, consistent with the effects of GLP-2 on normalizing barrier function, there was
a trend towards a decrease in claudin-7 expression with GLP-2 treatment in control animals
(Figure 3.10). Expression levels of the junction proteins, claudins-3, -15, occludin and ZO-1,
were unaffected by GLP-2 treatment. In comparison to control mice, KO mice demonstrated
elevated claudin-3 (p<0.05) and reduced claudin-7 (p<0.05). Furthermore, GLP-2 treatment
in KO mice had no effect on expression of any of these proteins. Finally, GLP-2 did not
affect localization of the tight junction proteins examined (Figure 3.11). Nonetheless,
consistent with healthy mice findings, IE-IGF-1R deletion-induced intracellular occludin was
also detected in this model of enteritis.
56
57
Figure 3.10 Protein expression of tight junctions in irinotecan-induced enteritis.
(A) Claudin (Cl)-3 (n=5-6), (B) Cl-7 (n=4-6), (C) Cl-15 (n=4-6), (D) occludin (n=4-6), and
(E) ZO-1 (n=4-6) expression levels were quantified by immunoblotting (*p<0.05). Vehicle
and GLP-2 treatments are presented by white and black bars, respectively.
58
59
Figure 3.11 Localization of tight junctions in irinotecan-induced enteritis.
Tight junctions were immunostained in red, the brushborder marker sucrase in green, and
nuclei in blue (n=3). Pictures subjected to comparison were captured under the same
exposure settings.
60
4 DISCUSSION
The intestinal epithelium is a selectively permeable barrier that restricts passage of harmful
substances while permitting absorption of nutrients. Trophic factors are known to enhance or
restore intestinal barrier function, and these include GLP-2, IGF-1, EGF and transforming
growth factor (TGF)-β1 (Benjamin et al., 2000; Peterson et al., 1996; Dignass and Podolsky,
1993; Hirano et al., 1995; Planchon et al., 1994). GLP-2 is unique because its actions are
largely intestinal-specific. Given that Teduglutide (human Gly2GLP-2) has recently been
approved for SBS, clinical use of GLP-2 has been demonstrated to be safe thus far; yet, a
large knowledge gap remains in our understanding of its mechanism of action. Therefore,
delineating signaling pathways of GLP-2 is imperative to identify possible side effects as
well as discovering potential novel clinical applications of this approved and intestinal-
specific therapeutic. Although some progress with respect to knowledge of GLP-2-mediated
intestinal growth has been established, the mechanism by which GLP-2 enhances barrier
function is poorly understood. Limited evidence has suggested that GLP-2 may affect tight
junctional proteins within the epithelium (Cani et al., 2009), but as the intestinal epithelial
cell does not express the GLP-2R (Yusta et al., 2000), this pathway is likely to involve a
downstream mediator. The current study reveals that IE-IGF-1R signaling is downstream of
GLP-2 in mediating barrier effects in states of health and enteritis.
The present study extends previous findings on the effects of GLP-2 in enhancing
intestinal barrier function. The in vivo effects of GLP-2 on lowering intestinal permeability
have been demonstrated in the context of low-grade inflammation associated with obesity,
61
using the leptin-deficient (ob/ob) mouse model (Cani et al., 2009). Within a cohort of ob/ob
mice, which included both untreated and prebiotic-treated animals, Cani et al. found that
higher circulating GLP-2 levels, suggested to be a response to prebiotic treatment, are
correlated with lower intestinal permeability, as measured by plasma levels of microbiota-
derived lipopolysaccharide (LPS) and of orally-administered FD4. Additionally, GLP-2
antagonist treatment abolishes the effects of prebiotics in lowering permeability to LPS and
reduced inflammatory tone, as determined by inflammatory cytokine levels (Cani et al.,
2009). Delzenne’s group further suggests a direct link between GLP-2 and intestinal
permeability by treating ob/ob mice chronically with GLP-2, which also results in reduced
LPS permeability and inflammatory tone. The present study extends previous findings by
demonstrating that exogenous GLP-2 reduced intestinal permeability to FD4 in vivo in
healthy, non-obese, animals. The macromolecule FD4 is a paracellular leakage marker, but is
not restricted to a particular segment of the GI tract. Thus, I also measured barrier function
using Ussing chambers in jejunal tissue, where the GLP-2R is the most abundant. In IGF-
1Rflox/flox
control mice bred on a C57BL/6 background, GLP-2 administration increased
jejunal resistance, indicative of reduced permeability to small ions. This is consistent with a
previous report on CD-1 mice that respond to GLP-2 treatment with reduced conductance,
the inverse of resistance (Benjamin et al., 2000).
Intestinal paracellular permeability is regulated by the epithelium, yet the IE cells
lack the GLP-2R (Yusta et al., 2000). The current study identifies, for the first time, the IE-
IGF-1R as a mediator of GLP-2-enhanced barrier function in vivo and ex vivo; in IE-IGF-1R
KO mice, GLP-2-lowered permeability to FD4 was abolished, and elevated jejunal resistance
in response to GLP-2 treatment was also reduced. Given that the IE-IGF-1R has previously
62
been demonstrated to mediate the growth effects of GLP-2 (Rowland et al., 2011), the
present findings extend and highlight the importance of IE-IGF-1R signaling in mediating
pleiotrophic intestinal effects of exogenous GLP-2.
Despite the finding that that enhanced barrier function in response to exogenous GLP-
2 required the presence of the IE-IGF-1R, vehicle-treated control and KO animals did not
differ in GI permeability to FD4. This indicates either endogenous GLP-2 play a negligible
role in modulating barrier function, or that the barrier effects of endogenous GLP-2 does not
require the IE-IGF-1R. This can be reconciled with studies using the GLP-2R-deficient
mouse model, in which jejunal resistance was not different between wild type and GLP-2R
null animals (Lee et al., 2012). Jejunal resistance of vehicle-treated mice was also not
significantly different between IGF-1Rflox/flox
control and IE-IGF-1R KO mice, although
there appeared to be a trend towards higher resistance in KO animals. There exists the
possibility of compensatory effects to have occurred in the IE-IGF-1R KO mice through up-
regulation of other growth factors known to improve barrier function (EGF, TGF-β1). It may
then be argued that GLP-2-treated IE-IGF-1R KO animals had already reached maximal
resistance. Further experiments using higher doses of GLP-2 can be performed to test
whether a greater resistance response could be triggered.
IGF-1R was deleted in IE cells by tamoxifen-activated Cre recombinase driven by the
villin promoter. Recombination induced by villin-Cre has been shown to persist in the
intestine for 60 days after tamoxifen withdrawal (el Heuval et al., 2004). Therefore, all
experimental timelines in the current study were conducted within the duration of 60 days.
The Brubaker lab has previously validated the inducible IE-IGF-IR-deficient mouse model
63
over a 1 day to 2 week time period, with receptor deletion achieved at 85% at the genomic
level and 89% at the mRNA transcript level (Rowland et al., 2011). For the present project,
functional confirmation of IE-IGF-1R deletion was done by measuring crypt-villus height,
which is known to increase with GLP-2 treatment, but is attenuated in the absence of the IE-
IGF-1R (Rowland et al., 2011). Several reports raise concern over the Cre/lox methodology.
For instance, tamoxifen toxicity has been shown in gastric epithelial, but not intestinal
epithelial cells (Huh et al., 2010). Moreover, Cre expression in the pancreatic β cell is known
to alter cell function (Lee et al., 2006; Pomplun et al., 2007). To control for potential
confounding experimental outcomes of tamoxifen, both control and KO animals were thus
treated with tamoxifen. However, future experiments should include villin-CreERT2+/0
animals as additional control for potential effects of Cre expression in the IE cells.
The paracellular transport route is primarily regulated by the apically situated tight
junctional complex. Electron micrographs of enterocyte ultrastructure did not show changes
in the tight junctional network between the four groups of animals. Similarly, Benjamin et al.
also did not detect any tight junction changes in response to GLP-2 treatment by electron
microscopy (Benjamin et al., 2000). Future studies could use freeze-fracture microscopy to
examine and quantify the length of tight junction strands. Nonetheless, GLP-2 has been
suggested, in the ob/ob mouse model, to upregulate tight junctional protein levels (occludin
and ZO-1) measured by a non-quantitative method of immunofluorescence image scoring
(Cani et al., 2009). Thus, we investigated selective tight junctional proteins, including ZO-1
and occludin. Additionally, we also selected the three proteins from the claudin superfamily
that were most highly expressed in the jejunum (Holmes et al., 2006). Claudin-3 and -7 are
considered barrier-forming sealing tight junction proteins (Van Itallie and Anderson, 2006).
64
The current findings show that GLP-2 increased protein expression of claudin-3 and -7,
without altering distribution, and these effects required the IE-IGF-1R. This suggests that
IGF-1R signaling in the IE cells triggers upregulation of claudin-3 and -7 at the protein level,
which ultimately reduces paracellular permeability. In contrast, pore-forming claudin-15 and
scaffolding ZO-1 were unaltered in terms of expression and localization.
Occludin is another transmembrane tight junctional protein, known for its sealing
effects (Cummins, 2012). Occludin is also known to be endocytosed and localized to
intracellular domains under conditions of barrier disruption (Cummins, 2012). In the current
studies, occludin expression was not changed by GLP-2 treatment or IE-IGF-1R disruption.
However, occludin localization in control mice was found to be highly concentrated along
the apical enterocyte membrane whereas, in the absence of the IE-IGF-1R, occludin shifted
to reside largely within the intracellular domain. Moreover, future studies to investigate other
tight junctional proteins will be important to map out potential signaling pathways in
enhancing intestinal barrier function. Taken together, the collective results in healthy animals
show that GLP-2 improved intestinal barrier function and required an IE-IGF-1R-dependent
pathway in altering tight junctional dynamics.
An enteritis model was utilized to interrogate the role of GLP-2 on barrier function
under the condition of inflammation. Irinotecan is anti-proliferative through inhibition of
topoisomerase I, an essential enzyme for a variety of DNA-associated processes including
replication, transcription, and recombination (Rivory and Robert, 1995). Irinotecan is a pro-
drug transformed into the much more active 7-ethyl-10-hydroxy-camptothecin by
carboxylesterases, subsequently glucuronized in the liver and released into the intestinal
65
lumen via the bile duct (Kawato et al., 1991). Tumour regression outcomes with irinotecan
are thus accompanied by dose-limiting gastrointestinal toxicity, manifested as mucositis and
diarrhea (Sears et al., 1999).
Unexpected, irinotecan treatment in control animals resulted in lowered GI
permeability to FD4 as well as increased jejunal resistance. The dose of irinotecan
administered was moderate and no diarrhea was detected nor was there mortality in the
experimental animals. Notwithstanding, these barrier changes were accompanied by
abnormal mucosal morphology and reduced crypt-villus growth. Precedent literature exists
for increased small intestinal resistance in response to irinotecan-induced mucositis in the rat
(Nakao et al., 2012). In addition, in a murine IL-2 KO model of colitis, colonic transmural
resistance is increased due to submucosal edema and inflammatory infiltration rather than
due to enhanced barrier function (Barmeyer et al., 2002). Therefore, the increased jejunal
resistance observed in irinotecan-treated animals may not be indicative of increased barrier
function. Interestingly, GLP-2 treatment restored crypt-villus growth, as well as GI
permeability and jejunal resistance in control animals that received irinotecan. This suggests
that the effects of GLP-2 under inflamed conditions results in normalization, instead of
enhancement, of barrier function, and these effects may be secondary to reduced mucosal
inflammation and reduced edema. Nonetheless, in irinotecan-induced enteritis, GLP-2-
normalized barrier function was dependent on the IE-IGF-1R as shown by reduced in vivo
permeability to FD4 and ex vivo jejunal resistance.
In irinotecan-induced enteritis, the effects of GLP-2-normalized barrier function may
have been independent of tight junctions, as the proteins examined by immunoblotting and
66
immunofluorescence, claudin -3, -7, -15, occludin and ZO-1, were unaltered by GLP-2
treatment. Nonetheless, there was a trend toward down-regulation of claudin-7 in response to
GLP-2 treatment, which appeared consistent with effects on barrier function. Given that
GLP-2 has been shown to attenuate irinotecan-induced intestinal mucositis in association
with reduced enterocyte apoptosis, the barrier effects of GLP-2 in this enteritis model may be
secondary to reduced inflammation and restored mucosal growth (Boushey et al., 2001).
Further investigations should also determine the resistance of the isolated epithelial layer, as
opposed to full thickness, and enteritis severity by scoring of H&E-stained jejunal tissue. In
addition, changes in colonic barrier function were not examined in the present studies and
may have contributed to altered GI physiology.
Further experiments are required to validate the specificity of the antibodies used
against the tight junction proteins examined. Possible experiments include staining using
negative control tissues, such as tissue lacking the particular protein or tissue from the
corresponding KO animal. In addition, antibody pre-absorption is an alternative technique.
Moreover, immunogold labeling of tight junction proteins could be utilized to determine
localization of the junctions more precisely than immunofluorescence.
One limitation to my studies is that potential alterations in the microbiome by IE-
IGF-1R deletion may confound results on intestinal permeability. GLP-2R KO mice exhibit
marked changes in the proportion of major bacterial microbiome constituents (Lee et al.,
2012). Thus, there remains a possibility that the effects of GLP-2 on the microbiome are
mediated by the IE-IGF-1R through epithelial-microbiota interactions. Another limitation is
that, although IE-IGF-1R was knocked out, signaling of IGF-1 and -2 could still occur via
67
binding to the insulin receptor (IR) on the epithelium. Whether IR activity affects intestinal
barrier function and whether binding of the IGFs to IR in the intestinal epithelium is
sufficient to induce barrier changes remain to be elucidated. Although the possibility of IR
altering permeability downstream of the IGFs cannot be ruled out, my results demonstrate
that inducible deletion of the IE-IGF-1R reduced the effects of GLP-2 in the enhancement of
barrier function in health.
In conclusion, the collective findings from these studies show that GLP-2 modulates
intestinal barrier function under both normal and inflamed conditions. The proliferative
effects of GLP-2 have been shown to require the IE-IGF-1R (Rowland et al., 2011), and the
present study extends these findings to demonstrate, for the first time, that the effects of
GLP-2 on barrier function also require the IE-IGF-1R. Furthermore, the expression of two
epithelial junctional proteins, claudin-3 and -7, were upregulated with GLP-2-enhanced
barrier function in the healthy mouse, and these effects also occurred in an IE-IGF-1R-
dependent fashion. Finally, under conditions of enteritis, GLP-2 improved intestinal
morphology and normalized intestinal permeability, actions that also required the IE-IGF-1R.
Taken together, I have shown that, in addition to promoting mucosal growth, changes in
intestinal barrier function mediated by GLP-2 signal through the IE-IGF-1R in health and
disease. Thus, these findings contribute to a mechanistic advancement in our understanding
of the intestinal-specific actions of GLP-2.
68
5 APPENDIX
5.1 Intestinal permeability and diabetes mellitus
Links have been established between diabetes mellitus (both type 1 and type 2) and
dysregulated intestinal barrier function. Increased intestinal permeability in diabetic patients
was first reported by Mooradian et al. in 1986 (Mooradian et al., 1986), followed by other
independent studies (Kuitunen et al., 2002; Sapone et al., 2006). Experimental and clinical
findings suggest that intestinal barrier defects play a role in type 1 diabetes pathogenesis;
prediabetic patients exhibit higher increase in intestinal permeability, and it is believed that
increased exposure to luminal antigens can lead to autoimmune destruction of β cells (Bosi et
al., 2006). Moreover, streptozotocin (STZ)-induced and non-obese diabetic mice have been
shown to exhibit increased bacterial translocation (Berg, 1985), and increased intestinal
conductance (Hadjiyanni et al., 2009), respectively. In addition, a series of recent studies has
emphasized the link between the composition of the intestinal microbiota, changes in gut
leakiness and the development of low-grade inflammation, leading to insulin resistance and
glucose-intolerance in animal models of type 2 diabetes (Cani et al., 2009; Brubaker and
Drucker, 2004; Delzenne and Cani, 2011).
5.2 Material and Methods
Both IGF-1Rfl/fl
control and IE-IGF-1R KO mice were induced with tamoxifen (1
mg/day) injections for 5 days. Subsequently, control and KO mice were treated with either
vehicle (0.1 mM citrate buffer) or a low dose of STZ (50 µg/g body weight) after fasting for
69
6 hr. Body weight and basal blood glucose levels were measured periodically throughout the
studies. Tail vein blood samples for glucose testing were measured using the OneTouch
Basic glucose meter (Lifescan Canada, Burnaby, BC). After overnight fasting, oral glucose
tolerance tests (OGTT) were conducted on the day 32 after the initial STZ injection, in which
mice were gavaged with glucose (1.5mg/g). Animals were fasted for 6 hr for insulin
tolerance tests (ITT) on day 34, and were injected (i.p.) with human biosynthetic insulin
(0.5U/kg; Novo Nordisk Pharmaceutical Industries, Toronto, ON). For both OGTT and ITT,
blood glucose was measured at t = 0, 10, 20, 30, 60, 90 and 120 min. FD4 permeability was
measured on day 36, as described in Section 2.2.
5.3 Summary of STZ studies
STZ was utilized to induce diabetes as a model of impaired barrier function.
Unexpectedly, these animals bred on a C56BL/6 background appeared resistant to low dose
of STZ administered, and developed only impaired glucose tolerance (Figure. 5.1).
Following STZ treatment, the IE-IGF-1R KO mice did not respond with differences in body
weight, susceptibility to diabetes onset, or altered glycemic control as compared to STZ-
treated controls (Figure 5.1A-D). STZ did not trigger elevated intestinal permeability, which
was also not different between the control and KO animals (Figure. 5.1E). Therefore, this
model was not pursued further.
70
71
Figure 5.1 STZ did not induce diabetes nor impaired barrier function
(A) Body weight and (B) blood glucose levels were monitored periodically through the
studies. (C) Oral glucose and (D) insulin tolerance were tested (n=3-8). Control and KO mice
are represented by black and red lines, respectively. Solid and dashed lines represent vehicle
and STZ treatements, respectively. (E) GI permeability to FD4 in streptozotocin-treated
control and KO mice (n=7-14).
72
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