intestinal cholecystokinin controls …...cholecystokinin (cck) is a gut peptide involved in the...
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INTESTINAL CHOLECYSTOKININ CONTROLS GLUCOSE PRODUCTION THROUGH A NEURONAL NETWORK
by
Grace Wing Chee Cheung
A thesis submitted in conformity with the requirements for the degree of MASTER OF SCIENCE
DEPARTMENT OF PHYSIOLOGY UNIVERSITY OF TORONTO
© Grace Wing Chee Cheung Copyright 2010
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Title of thesis: Intestinal cholecystokinin controls glucose production through a neuronal
network
Name of candidate: Grace Wing Chee Cheung
Degree: Master of Science
Department of Physiology, University of Toronto
Year of convocation: 2010
GENERAL ABSTRACT
Cholecystokinin (CCK) is a gut peptide involved in the regulation of energy homeostasis by
duodenal lipids via a neuronal network. However, it is unknown whether CCK also regulates
glucose homeostasis through a neuronal network. Using an in vivo rat model, we demonstrated
that duodenal CCK-8 (biologically active form of CCK) can lower glucose production through
the activation of a gut-brain-liver axis via CCK-A receptors, and this glucose-regulatory effect is
physiologically relevant. Since duodenal lipids can also lower glucose production through a gut-
brain-liver axis, we verified that this duodenal-lipid effect is mediated by CCK-A receptor
activation. Lastly, in rats fed on a high-fat diet for three days, duodenal CCK failed to suppress
glucose production, suggesting a state of CCK-resistance. In summary, these findings revealed
that intestinal CCK can regulate glucose homeostasis through a neuronal network and suggest
that intestinal CCK resistance may contribute to hyperglycemia in response to high-fat feeding.
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Acknowledgement
First of all, I would like to express my deepest gratitude to my supervisor, Dr. Tony Lam. He has
given me opportunities to develop in various ways academically. He taught me how to design
experiments, how to write scientific papers, how to present my findings. I am also thankful for
his willingness in guiding my life decisions. Moreover, I would like to thank my supervisory
committee members, Dr. Harvey Anderson and Dr. Adria Giacca. I am grateful to have such
wonderful scientists in my supervisory committee. Dr. Anderson and Dr. Giacca taught me to not
be satisfied with a single answer but to be curious and seek for other explanations. In addition to
the supervisory figures, the laboratory had been an integral part of my graduate experience and it
would not have been so rewarding without these people: Carol Lam, Madhu Chari, Andrea
Kokorovic, Clair Yang, Penny Wang and Teresa Lai.
Aside from the academic support, I would also like to thank the following people. First, I would
like to thank my parents, Edward Cheung and Enid Li, for their unconditional support and
encouragement to pursue my interests. My sister, Bonnie Cheung, for all the joy she has brought
to my life. My significant other, Terry Wong, for standing by my side through all the ups and
downs and providing all the support I ever needed. Last but not least, I would like to thank my
Lord for His continual guidance and all the wonderful plans He made for me, I will continue to
trust Him with all my heart for all the challenges ahead.
“Trust in the Lord with all your heart and lean not on your own understanding; in all
your ways acknowledge him, and he will make your paths straight.” Proverbs 3:5
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Table of Contents
Acknowledgement ......................................................................................................................... iii
Table of Contents ........................................................................................................................... iv
List of Figures ................................................................................................................................. v
List of Tables ................................................................................................................................. vi
List of Abbreviations .................................................................................................................... vii
Publication that contributed to this thesis ...................................................................................... ix
1 Introduction .................................................................................................................................. 1
1.1 Obesity and Diabetes .................................................................................................... 1
1.2 Regulation of Glucose Homeostasis: The Role of the Small Intestine ......................... 3
1.3 Gastrointestinal Peptides ............................................................................................... 6
1.3.1 Gastric Peptide ............................................................................................... 6
1.3.2 Upper Intestinal Peptide ............................................................................... 10
1.3.3 Lower Intestinal Peptide .............................................................................. 15
1.4 Neuronal Regulation of Glucose Homeostasis by the Small Intestine ....................... 22
1.4.1 Hepatoportal Glucose Sensor ....................................................................... 23
1.4.2 Lipid-induced gut-brain-liver axis ............................................................... 25
2 General Hypothesis and Aims ................................................................................................... 29
3 General Materials and Methods ................................................................................................. 35
4 Results ........................................................................................................................................ 50
5 Discussion .................................................................................................................................. 72
6 Future Directions ....................................................................................................................... 78
7 Conclusion ................................................................................................................................. 81
8 References .................................................................................................................................. 82
v
List of Figures
Figure 1 Upper intestinal lipids activate a gut-brain-liver axis to suppress glucose production ......................................................................................................................28
Figure 2 Working hypothesis – upper intestinal lipids stimulate CCK/CCK-A receptor signaling to suppress glucose production through a gut-brain-liver axis ......................34
Figure 3 Schematic representation of the working hypothesis – duodenal CCK activates CCK-A receptors to regulate glucose production and experimental design (pharmacological approach). .........................................................................................58
Figure 4 Duodenal CCK activates CCK-A receptors to suppress glucose production ................59
Figure 5 Schematic representation of the working hypothesis – duodenal CCK activates CCK-A receptors to regulate glucose production and experimental design (molecular approach). ....................................................................................................60
Figure 6 Duodenal CCK suppresses glucose production in LETO but not in CCK-A receptor deficient OLETF rats .......................................................................................61
Figure 7 Schematic representation of the working hypothesis – duodenal CCK activates a gut-brain-liver axis to suppress glucose production and experimental design ..............62
Figure 8 Duodenal CCK can activate a gut-brain-liver axis to regulate glucose production ......63
Figure 9 Schematic representation of the working hypothesis – duodenal CCK signaling is downstream of lipid-sensing to regulate glucose production and experimental design .............................................................................................................................64
Figure 10 Duodenal CCK-A receptor activation is required for lipids to lower glucose production ......................................................................................................................65
Figure 11 Pharmacological inhibition of CCK-A receptors in the gut disrupts glucose homeostasis during refeeding ........................................................................................66
Figure 12 Molecular inhibition of CCK-A receptors in the gut disrupts glucose homeostasis during refeeding ............................................................................................................67
Figure 13 Schematic representation of the working hypothesis – duodenal CCK fails to suppress glucose production in response to high fat feeding and experimental design .............................................................................................................................68
Figure 14 Duodenal CCK fails to suppress glucose production following 3-days of high-fat feeding ...........................................................................................................................69
vi
List of Tables
Table 1 Dietary contents of the regular chow and the lard-oil enriched high fat diet. .................49
Table 2 Plasma insulin and glucose concentrations for SAL and CCK8 during basal and clamp conditions .........................................................................................................................70
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List of Abbreviations AA Arachidonic acid
AMPK Adenosine monophosphate kinase
ANOVA Analysis of variances
ATP Adenosine triphosphate
B Bound fraction
B0 Total binding
cAMP cyclic adenosine monophosphate
CCK Cholecystokinin
CCK-A receptor Cholecystokinin-A receptor
CCK-B receptor Cholecystokinin-B receptor
GHS-R Growth hormone secretagogue receptor
GIP Glucose-dependent insulinotropic polypeptide
GIP-R Glucose-dependent insulinotropic polypeptide receptor
GLP-1 Glucagon-like peptide-1
GLP-1R Glucagon-like peptide-1 receptor
GLUT4 Glucose transporter 4
GPR40 G-protein coupled-receptor 40
GSIS Glucose-stimulated insulin secretions
HFD High-fat diet
ICV Intracerebroventricular
IP Intraperitoneal
IP3 Inositol triphosphate
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IV Intravenous
KATP channels ATP-sensitive potassium channels
Kv channels voltage-dependent potassium channels
LCFA Long chain fatty acid
LETO rat Long-Evans Tokushima Otsuka rat
NMDA receptor N-methyl-d-aspartate receptor
NTS Nucleus of the solitary tract
OLETF rat Otsuka Long-Evans Tokushima Fatty rat
OXM Oxyntomodulin
PKA Protein kinase A
PKC Protein kinase C
PLA2 Phospholipase A2
PLC Phospholipase C
PYY Peptide YY
Ra Rate of appearance
Rd Rate of disappearance
RIA Radioimmunoassay
SC Standard chow
SD rat Sprague Dawley rat
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Publication that contributed to this thesis
- Cheung GW
, Kokorovic A, Lam CK, Chari M and Lam TK. Intestinal cholecystokinin
controls glucose production through a neuronal network. Cell Metab. 10: 99-109, 2009.
1
1 Introduction
1.1 Obesity and Diabetes
According to the World Health Organization, approximately 1.6 billion adults (age 15+; body
mass index ≥ 25 and <30) were overweight, with at least 400 million adults obese (BMI ≥ 30) in
2005. Obesity is associated with type 2 diabetes, cardiovascular disease and various cancers
[1;2]. It has been proposed that elevated plasma free fatty acids may serve as a causative link
between obesity, insulin resistance and diabetes [3;4]. While diabetes is a disease characterized
by hyperglycemia due to impairments in the regulation of glucose homeostasis, it has been
demonstrated that free fatty acids can induce insulin resistance [5], strongly suggesting a link
between obesity and type 2 diabetes.
In 2000, approximately 170 million individuals were affected by diabetes, with the affected
population expected to double by 2030 [6]. There are two forms of diabetes mellitus: type 1 and
type 2. Type 1 diabetes is an autoimmune disorder in which there is near total deficiency in
insulin secretions due to the destruction of insulin-producing β-cells [7]. On the other hand, type
2 diabetes is the more common form, affecting more than 90% of the patients with diabetes, and
it is linked to a combination of insulin resistance and deficient insulin secretion [8]. Insulin is
responsible for (i) stimulating glucose uptake into muscle and fat tissues (ii) suppressing glucose
production in the liver [8]. Hence, insulin resistance can lead to excessive glucose in the
circulation resulting in chronic hyperglycemia. Various complications can result from chronic
hyperglycemia [9;10], including retinopathy [11], kidney failure [12], increased risks of
cardiovascular disease [13] and stroke [9]. As such, it is essential to dissect the mechanisms
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involved in the regulation of glucose homeostasis and identify possible interventions to restore
the regulation.
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1.2 Regulation of Glucose Homeostasis: The Role of the Small Intestine
The regulation of glucose homeostasis can be distinguished in two different states. In the fasting
basal state, plasma glucose levels are derived endogenously. There are primarily two hormones
involved in the regulation of glucose homeostasis: glucagon and insulin [14]. Glucagon is
secreted from pancreatic α-cells to prevent hypoglycemia through stimulating hepatic glucose
production [15]. In contrast, insulin prevents hyperglycemia through stimulating glucose uptake
and suppressing glucose production [8]. In the fasting state, glucagon secretion rises while
insulin secretion falls to ensure that there is sufficient glucose circulating.
Upon meal intake, glucose regulation changes from the fasting state to the fed state. In the fed
state, there are both exogenous and endogenous sources of glucose. Exogenous glucose is
absorbed from the gastrointestinal tract as nutrients become available for absorption in the small
intestine. Hence, glucose absorption rate depends on the rate at which nutrients are emptied from
the stomach to the small intestine, known as the gastric emptying rate [16]. As exogenous
glucose arrives in the circulation, a biphasic insulin secretion response is stimulated. In response
to an intravenous (iv) glucose challenge, insulin is rapidly secreted to reach an initial peak in 5 to
7 minutes [17], and this early-phase of insulin release lasts about 10 to 15 minutes. On the other
hand, the late-phase is characterized by a more sustained insulin secretion lasting several hours
[17]. In addition to the responses in insulin, glucagon secretion is suppressed. Together,
endogenous glucose production is suppressed, while glucose uptake is augmented to allow
plasma glucose concentrations to return to baseline values such that the fasting state resumes.
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Since the small intestine is one of the first sites of contact of the body to incoming nutrients, the
small intestine is at a strategic location to exert actions on the regulation of glucose homeostasis
during feeding. The regulation of glucose homeostasis by the small intestine can largely be
distinguished into two types of mechanisms: (i) indirect actions through gastrointestinal peptides
(ii) direct neuronal pathway.
For the indirect actions mediated by gastrointestinal peptides, the effects are mainly exerted
through modulating the gastric emptying rate and pancreatic hormone levels in response to the
entry of nutrients as will be described in more details later. Through the modulation of gastric
emptying rate, the rate at which glucose is delivered to the small intestine can be controlled to
affect the rise of glucose in the blood [16]. On the other hand, the secretion of insulin and
glucagon can also be potentiated or inhibited by gastrointestinal peptides to regulate plasma
glucose levels differentially.
For the neuronal regulation of glucose homeostasis by the small intestine, a hepatoportal sensor
[18;19] and a gut-brain-liver [20] neuronal axis induced by glucose and lipid respectively have
been identified. While it seems that the mechanisms of action for the gastrointestinal peptides are
unrelated to the neuronal mechanisms, gastrointestinal peptides have been shown to regulate
food intake through neuronal actions [21]. Hence, it is possible that gastrointestinal peptides are
involved in the activation of the neuronal networks in the regulation of glucose homeostasis.
In the following section, we will first focus on the gastrointestinal peptides that are modulated by
duodenal nutrients (duodenum is the first segment of the small intestine) and the role of each in
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the regulation of glucose homeostasis, and where applicable, the neuronal network responsible
for modulating food intake by each gastrointestinal peptide. The gastrointestinal peptides will be
presented according to their anatomical origin.
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1.3 Gastrointestinal Peptides 1.3.1 Gastric Peptide
1.3.1.1 Ghrelin Ghrelin was isolated in 1999 as an acylated 28-amino acid peptide with the acylation required for
its bioactivity [22]. Ghrelin is predominantly secreted in the stomach and duodenum [22-25] and
its levels are usually increased prior to eating, while reduced in response to nutrient entry
[26;27]. In particular, intraduodenal administration of glucose [28-31], amino acids [31], or
lipids [31-34] can all suppress ghrelin secretion to discontinue the fasting signal. With respect to
the regulation of secretion by lipids, lipids in the form of triglycerides must first be broken down
by lipases to release fatty acids [33]. Following the release of fatty acids, the accumulation of
long chain fatty acids (LCFA) [34] in the duodenum leads to the secretion of another
gastrointestinal peptide called cholecystokinin (CCK) and the binding of CCK to cholecystokinin
A (CCK-A) receptors is necessary for lipids to suppress ghrelin secretion [32].
Since ghrelin is mostly secreted during the fasting state, it is not surprising that ghrelin plays a
significant role in the maintenance of fasting blood glucose levels through modulating insulin
secretion. In humans, ghrelin increases glucose while decreasing plasma insulin levels [35].
Ghrelin is a natural ligand for the growth hormone secretagogue receptor (GHS-R) [22;36;37],
the blockade of GHS-R resulted in reduced fasting blood glucose concentrations [38] and
increased plasma insulin levels [39]. This suggests that ghrelin plays a significant role in the
regulation of fasting blood glucose through playing an inhibitory role on insulin secretion.
Interestingly, the source of ghrelin for the effects on insulin may not originate from the stomach
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or duodenum [40], but from the pancreas [38]. After undergoing gastrectomy to eliminate the
secretion of ghrelin by the stomach, it was found that the administration of ghrelin-antagonist
could still augment glucose-stimulated insulin secretions (GSIS), strongly suggesting that a local
source of ghrelin in the pancreas is responsible for the GSIS effects. However, the regulation of
secretion for pancreatic ghrelin remains to be determined. Nonetheless, GHS-R have been
located in pancreatic islets [25;38;41-44], suggesting that GHS-R in the pancreas may mediate
the effects of ghrelin on pancreatic insulin secretions. Indeed, in isolated rat islets of GHSR-
knockout mice, GSIS was found to be significantly greater than that of control mice [39],
supporting the notion that endogenous ghrelin acts on the GHS-R of pancreatic islets to regulate
GSIS. At the level of the islets, ghrelin failed to suppress insulin secretion in the presence of
pertussis toxin or voltage-dependent potassium (Kv) channel blockers [39], suggesting that
ghrelin can activate a specific isoform of G-proteins, which can lead to the activation of Kv
channels. In particular, it was confirmed with a molecular antisense approach that the isoform of
G-protein involved is the G-alpha i2 form [39]. Importantly, when the activity of Kv channels
was prevented, the rise in intracellular calcium levels associated with insulin release was
attenuated [39]. On the basis of these findings, ghrelin binds to the GHS-R on the islets to
decrease the activity of glucose-stimulated action potentials of the islet membranes marked by
decreased frequency and amplitude of firings [39]. As the activity of action potentials reduces,
the duration of the bursting potentials stimulated by glucose is shortened, preventing further
calcium-dependent insulin exocytosis. In turn, the suppression of GSIS allows the body to
restrain insulin secretion in the fasting state whereas the restraining effect on insulin can be lifted
in the presence of duodenal nutrients through the inhibition of ghrelin. Conversely, ghrelin does
not affect glucagon secretion [45].
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In addition to its effects on insulin secretion, ghrelin may also affect insulin sensitivity. In
humans, ghrelin infusion induced acute insulin resistance [46]. In another study, it was shown
that ghrelin may induce insulin resistance specifically in muscle to reduce glucose disposal while
having no effects on glucose production [47]. On the other hand, the ablation of ghrelin
improved peripheral insulin sensitivity in diabetic mice by affecting insulin action both on
glucose production and glucose disposal [48]. Although there are disagreements regarding the
tissue-specific effects of ghrelin, these studies in general suggest that ghrelin is responsible for
inducing insulin resistance. In contrast, there is also a study showing otherwise, that ghrelin may
improve peripheral insulin sensitivity while hampering hepatic insulin sensitivity [49]. Hence,
the role of ghrelin on insulin sensitivity remains controversial.
Nonetheless, it is commonly accepted that ghrelin can affect glucose homeostasis through its
action on gastric emptying. When ghrelin is infused, there is higher postprandial glucose as a
result of accelerated gastric emptying [50]. By increasing the gastric emptying rate, ghrelin
accelerates the speed at which glucose is delivered to the duodenum for absorption. There are
findings suggesting that ghrelin may accelerate gastric emptying through a neural mechanism.
First, GHS-R has been identified in nerves fibers belonging to the enteric nervous system [51]. In
rats, ghrelin enhanced contractions in strips of the gastric body while tetrodotoxin (neurotoxin)
abolished the effect, supporting that the intrinsic enteric nervous system may be involved [52]. In
addition to the enteric nervous system, in rats pretreated with capsaicin, atropine, or cervical
vagotomy, the ability of ghrelin to accelerate gastric emptying was abrogated [52]. Since there
are ghrelin receptors expressed in vagal afferent neurons [53], ghrelin may stimulate the vagus
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nerve directly or through the enteric nervous system to accelerate gastric emptying. The
increased gastric emptying rate can then allow nutrients to be delivered to the duodenum for
absorption as soon as they are available. As nutrients enter the duodenum, the nutrients can
stimulate a negative feedback system on gastric emptying by inhibiting ghrelin secretion,
resulting in the blunting of postprandial hyperglycemia.
As mentioned previously, ghrelin is mostly secreted in the fasting state, it is not surprising that
ghrelin can stimulate feeding in rats [53]. In response to ghrelin, the arcuate nucleus in the
hypothalamus is activated as indicated by fos-staining, and both the feeding and hypothalamic
activation responses are abolished in response to capsaicin treatment, gastric branch vagotomy,
or subdiaphragmatic vagotomy [53]. Hence, a neuronal network connecting the gut and the brain
should exist to mediate the effects of ghrelin on feeding, suggesting that there is a gut-brain
neuronal axis to allow communication between the gut and the brain.
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1.3.2 Upper Intestinal Peptide
1.3.2.1 Cholecystokinin CCK is a gut peptide that has been discovered by Ivy and Oldberg in 1928 to stimulate gall
bladder contractions [54]. In the intestine, CCK is synthesized as a 115 amino acid prepro-CCK
polypeptide. The prepro-CCK polypeptide then undergoes multiple posttranslational cleavages to
generate the shortest and the complete biologically active form of CCK-8 [55].
CCK is released basolaterally by I cells lining the mucosa of the duodenum to the surrounding
areas such as the bloodstream and nerve endings. Duodenal glucose, lipids, and proteins have all
been shown to stimulate CCK secretion [29;56-58]. For lipids, the release of LCFAs is essential
for the release of CCK [59-61]. While proteins can stimulate CCK secretions [62], individual
amino acids such as tryptophan [63] and phenylalanine [64] can also stimulate CCK release.
Given that CCK is released in response to nutrients, CCK serves many important functions
pertaining to the digestion and absorption of nutrients, including the stimulation of pancreatic
secretions, excretion of bile into the intestine, and slowing of gastric emptying [65].
In the context of glucose homeostasis regulation, CCK can act through two different mechanisms
to reduce postprandial hyperglycemia. First, there are mixed reports regarding the ability of CCK
to regulate insulin secretion in physiological setting. While some studies demonstrated that CCK
may potentiate GSIS [66-75], this effect was absent in other studies [76-80], questioning the
physiological relevance of this effect. Nonetheless, systemic administration of CCK induced a
biphasic response of insulin secretions [81], with an early rise after a minute, and a slow second
phase, associated with a concomitant drop in plasma glucose. First, CCK may potentiate GSIS in
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rat islets via interacting with the alpha q/11 subunit of G-proteins [82]. This subunit of G-protein
has been implicated in the phospholipase C (PLC) pathway, suggesting that CCK may potentiate
GSIS through the PLC system [82] to increase the production of inositol triphosphate (IP3) [82]
and intracellular calcium levels [83] to potentiate calcium-dependent insulin secretions. A
product of PLC activation is diacylglycerol (DAG), which is an activator of protein kinase C
(PKC). Indeed, CCK-8 has been shown to activate PKC through stimulating PLC [84].
Subsequent to PKC activation, CCK may activate phospholipase A2 (PLA2) to form arachidonic
acid (AA) [85] in isolated rat islets. It has been shown that the activation of PLA2 is partly
dependent on protein kinase C (PKC) and independent of calcium for the effects of CCK on
insulin [86]. Nonetheless, although CCK may potentiate GSIS through these mechanisms, these
effects may be pharmacological rather than physiological due to the high dosage used.
Secondly, a widely accepted mechanism by which CCK can regulate glucose homeostasis is
through the modulation of gastric emptying rate. CCK can reduce postprandial hyperglycemia
[87] by slowing the delivery of glucose to the duodenum in humans [80;88]. There are two types
of CCK receptors: CCK-A and CCK-B receptors, which reside in the periphery (predominantly
in the gastrointestinal system) and brain respectively [89]. For the regulation of gastric emptying
by CCK, CCK-A receptors are essential to stimulate the contraction of the pyloric sphincter [90-
95]. Interestingly, CCK may not need to bind to CCK-A receptors on the pyloric sphincter to
regulate gastric emptying [96]. Rather, CCK may activate CCK-A receptors on capsaicin-
sensitive vagal afferent C-fibers [97], which then regulate gastric emptying through the gastric
branch of the vagus nerve [98]. Hence, CCK regulates postprandial glucose homeostasis through
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the regulation of gastric emptying, and it may also potentiate GSIS when used
pharmacologically.
Moreover, CCK is the first gut peptide implicated in the control of food intake [99]. Specifically,
CCK has been shown to inhibit food intake in rats [100;101], humans [102], and mice [103]. The
effect of CCK on food intake is mediated by CCK-A receptors [104] and it is abolished in rats
subjected to abdominal vagotomy or gastric branch vagotomy [101], indicating that the vagus
nerve is required. Lastly, there is evidence that peripheral CCK can stimulate activation of
different brain areas via capsaicin-sensitive vagal afferent [105], strongly supporting that there is
direct communication between the gut and the brain.
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1.3.2.2 Glucose-dependent Insulinotropic Polypeptide Glucose-dependent insulinotropic polypeptide (GIP) is a 42-amino acid hormone [106]
expressed by K cells [107] with the highest concentration in the duodenum and jejunum in
humans [108]. GIP release can be induced by intraduodenal administration of glucose [58;109-
112], lipids [113-115], proteins [116] and amino acids [115]. For lipids in particular, long-chain
triglycerides are more potent than medium-chain triglyceride in stimulating GIP release in dogs
[115]. Interestingly, it has recently been found that free fatty acids can also stimulate the
secretion of GIP through a G-protein coupled receptor known as G-protein coupled-receptor 40
(GPR40) [117].
GIP is most commonly known as an “incretin”. Incretins are hormones that are responsible for
producing a greater insulin response to oral glucose than an equivalent amount of glucose
administered intravenously. Hence, one of the most important functions of GIP is its ability to
potentiate GSIS. It has been found that GIP has hypoglycemic effects through potentiating GSIS
in rats [118] and humans [119], with a specific effect on the early-phase insulin release [120]. In
studies using isolated rat pancreatic islets [121] and rat pancreas [122], it is suggested that GIP
acts directly on pancreatic islets to exert its potentiating effects. In the pancreas, the expression
of a receptor for GIP (GIP-R) has been identified [123], suggesting that GIP may activate the
GIP-R on pancreatic islets for its potentiating effect. Indeed, in GIP-R null mice, GIP failed to
augment GSIS [124]. Following the stimulation by GIP, there is increased cyclic adenosine
monophosphate (cAMP) production [123], and cAMP can lead to the activation of cAMP-
guanine nucleotide exchange factor (GEF) II (Epac2) and protein kinase A (PKA). Importantly,
treatment of pancreatic β-cells with either antisense oligodeoxynucleotides against Epac2 or the
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PKA antagonist H-89 demonstrated that both Epac2 and PKA are involved in the potentiating
effects of GIP [125]. Moreover, the closure of adenosine triphosphate (ATP)-sensitive potassium
(KATP) channels has also been found to be critical for the potentiation of insulin secretion by
cAMP [126] and GIP-induced potentiation of GSIS [127]. The closing of KATP channels alters
the membrane potential such that voltage-dependent calcium channels can be opened to increase
intracellular calcium levels. Indeed, the voltage-dependent calcium channels to allow the influx
of extracellular calcium [128] is essential as there is an absence of increases in intracellular
calcium levels in the presence of ethylene glycol tetraacetic acid to chelate extracellular calcium
[128]. In brief, GIP decreases KATP channel activity through the actions of PKA and Epac2 to
depolarize the membranes. As the membrane depolarizes, the opening of voltage-dependent
calcium channels allows the influx of calcium to increase calcium-dependent exocytosis of
insulin.
While it is clear that GIP potentiates GSIS, GIP appears to stimulate glucagon secretions in both
rats [118] and humans [129]. Since glucagon would exert an upregulating effect on circulating
glucose and GIP has been shown to lower glucose levels, it is unclear how this stimulatory effect
on glucagon influences the regulation of glucose by GIP.
Contrary to the hormones already discussed, GIP does not inhibit gastric emptying [130] in
humans. Hence, the main mechanism by which GIP regulates glucose homeostasis is through the
potentiation of GSIS.
GIP has no effects on food intake [131].
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1.3.3 Lower Intestinal Peptide
1.3.3.1 Glucagon-like peptide-1 Glucagon-like peptide-1 (GLP-1) is expressed in L cells [132] found in the ileum and colon of
rats, pigs and humans [133]. There are two active forms of GLP-1: GLP-17-37 and GLP-17-36-NH2
[134]. In humans and animal models, the secretion of GLP-1 demonstrates a biphasic profile
with an acute first peak of secretion within 15-30 minutes of nutrient ingestion and a second
sustained peak within 60-90 minutes [135]. However, nutrients normally do not reach the ileum
or colon within 15-30 minutes to account for the first peak of GLP-1 secretion [136], suggesting
that neural mechanisms originating from the proximal gut may play a role in the stimulation of
GLP-1.
Indeed, intraduodenal administration of glucose [58;137-139] and fat [140-144] can both
stimulate the release of GLP-1. With respect to duodenal glucose-stimulated GLP-1 release, the
concomitant transport of sodium ions and glucose via the sodium-glucose cotransporter-1 is
required while glucose metabolism is not necessary [138]. On the other hand, the mechanism(s)
by which duodenal fats stimulate GLP-1 release appears to be more complex. Upon entry of
lipids into the duodenum, the release of LCFAs through fat hydrolysis is essential to stimulate
GLP-1 [59;140]. Following the release of LCFAs, both CCK and GIP have been implicated in
mediating the stimulation of GLP-1 release. When the LCFA sodium oleate was coadministered
with a CCK-A receptor blocker, the rise of GLP-1 was abolished, indicating the importance of
CCK signaling in GLP-1 release in humans [140]. On the other hand, the involvement of GIP in
the stimulation of GLP-1 secretion may be species-dependent. In rats, GIP has been suggested to
mediate the stimulation of GLP-1 by duodenal fat [142], however, GIP had no effects on GLP-1
16
secretion in humans [145;146]. GIP can stimulate GLP-1 secretion through the hepatic branch of
the vagus nerve in rats, and stimulating the distal end of the celiac branch of the
subdiaphragmatic vagus nerve also results in the secretion of GLP-1 [144]. In light of these
findings, it is probable that a neural mechanism exists between the proximal and distal intestine
for duodenal nutrients to stimulate GLP-1 secretion. Two neurotransmitters are proposed to be
involved: gastrin-releasing peptide (GRP) [143] and acetylcholine [147]. In the presence of the
GRP antagonist, the GLP-1 response to duodenal fat was abolished [143]. Additionally,
cholinergic innervations are also important as atropine or the M1 muscarinic-receptor antagonist
pirenzepine completely abolished fat-induced GLP-1 secretion in rats [147]. Hence, a complex
network exists to mediate the release of GLP-1 in response to duodenal nutrients.
In addition to GIP, GLP-1 is also known for its function as an incretin to regulate postprandial
glucose levels [148] through its potentiating effects on GSIS. As GLP-1 is secreted, it can bind to
GLP-1 receptors on β-cells [149;150] to induce depolarization of pancreatic β-cells [151]. The
mechanism by which GLP-1 induces depolarization requires the closure of KATP channels [151].
With cAMP levels increased in response to GLP-1 receptor activation [152], both PKA and
Epac2 activation were found to be required to potentiate insulin secretion [125]. In particular,
PKA activation is required in mediating the closure of KATP channels [153;154]. Additionally,
other pathways such as a calmodulin-mediated pathway have also been implicated. In the
presence of the calmodulin inhibitor, the inhibitory actions of GLP-1 on the membrane KATP
conductance and the resultant membrane depolarization were completely reversed [155]. At the
level of the KATP channels, GLP-1 seems to potentiate ATP sensitivity to KATP channels such that
less ATP is required to close the KATP channels [151]. Subsequent to membrane depolarization,
17
the activation of L-type voltage-dependent calcium channels allows intracellular calcium to
increase [156;157] leading to calcium-dependent exocytosis of insulin. Finally, membrane
repolarization is controlled by the activity of voltage-dependent potassium (Kv) channels. In
particular, the inhibition of Kv channels has been shown to be dependent on both the PKA [152],
and the PKC-ζ pathways [158] to enhance (larger and prolonged) action potentials in the β-cells
[152]. Hence, GLP-1 can potentiate GSIS through modulating membrane potential via complex
mechanisms.
Moreover, GLP-1 has also been shown to suppress endogenous glucagon secretion, which may
contribute to its hypoglycemic effects [159-162]. Importantly, the suppressive effect of GLP-1
on glucagon secretion is glucose-dependent to prevent hypoglycemia from the lack of glucagon
[163]. Interestingly, GLP-1 receptors are not expressed in α-cells, meaning that GLP-1 should
regulate glucagon through an indirect mechanism [150]. Indeed, GLP-1 may act on the GLP-1
receptors on the δ-cells [149;150] to stimulate the secretion of somatostatin, then somatostatin
can act on the somatostatin receptor-2 to suppress glucagon secretion [164].
While GLP-1 can exert insulinotropic and glucagonostatic effects, GLP-1 can also regulate
glucose through altering the gastric emptying rate. Specifically, it has been shown that GLP-1
can inhibit gastric emptying to attenuate rises in glucose [127;165-167].
In summary, GLP-1 can regulate glucose homeostasis through acting on insulin and glucagon
secretions and gastric emptying. Interestingly, it has been found that GLP-1 can activate
neuronal mechanisms to increase insulin secretion.
18
Despite that GLP-1 can exert its incretin effects through direct interaction with β-cells, GLP-1 is
rapidly degraded by DPP-IV [168], thus it is unclear whether the short half-life of GLP-1 would
allow it to act on the pancreas directly. Hence, a rapid neural mechanism may actually be
responsible for the actions of GLP-1 on insulin secretion. As GLP-1 is released, it can enter the
portal vein to stimulate GLP-1 receptors expressed on nodose ganglions to improve glucose
disposal rate [169] through augmenting insulin secretion [169;170]. Importantly, when the
ganglionic blocker chlorisondamine is coinfused with GLP-1 into the portal vein, the insulin
secretion response is abolished [169]. Therefore, a neural mechanism is proposed to mediate the
portal GLP-1 effects on insulin secretion. Indeed, it has been found that administration of GLP-1
into the portal vein increased the firing rates of the hepatic afferent and pancreatic efferent
nerves [171], suggesting that a vagal hepatopancreatic reflex pathway may be involved in the
mediation of the insulinotropic effects of GLP-1 in addition to the endocrine pathway.
Moreover, GLP-1 can also activate a neuronal network to regulate food intake. In both humans
[172] and rats [173], GLP-1 has been shown to reduce food intake. Importantly, the satiety effect
is abolished in rats given subdiaphragmatic vagotomy or transection of a brainstem-
hypothalamic pathway [174]. These findings together suggest that GLP-1 can stimulate a gut-
brainstem-hypothalamic neuronal axis to regulate food intake, supporting the existence of a gut-
brain neuronal axis.
19
1.3.3.2 Oxyntomodulin Oxyntomodulin (OXM) is a 37-amino acid peptide expressed [175] throughout the
gastrointestinal tract [176]. OXM is expressed in L cells and the expression of OXM increases
from the duodenum to the ileum [176]. While OXM is mainly expressed in the lower intestine,
the release of OXM has only been directly assessed in a study which subjected received
intraduodenal administrations of a liquid meal or oleic acid [177]. Since OXM is also a product
of the proglucagon gene expressing GLP-1, OXM is believed to be co-secreted with GLP-1
[178].
Despite that OXM has been isolated and sequenced in 1994 [179], its action on glucose
homeostasis remains poorly studied. In response to glucose tolerance test, OXM administration
lowered blood glucose without affecting gastric emptying in mice [180], whereas OXM has been
shown to reduce gastric emptying rate in humans [181]. Although the effects of OXM on gastric
emptying rate remains to be clarified, it has been suggested that OXM can stimulate insulin
release in a glucose-dependent manner [180;182]. Interestingly, an oxyntomodulin receptor has
not been clearly demonstrated [183], whereas it has been shown that the action of OXM on
insulin secretion is dependent on GLP-1R [180]. In β-cells, cAMP formation is increased in
response to OXM [180], suggesting that OXM may bind to GLP-1 receptors on β-cells, and
potentiate GSIS through a cAMP-dependent mechanism.
Although OXM can reduce food intake [184], the mechanism of action remains largely
unknown.
20
1.3.3.3 Peptide YY Peptide YY (PYY) is synthesized in the body as a 36-amino acid peptide (PYY1-36) [185]. After
being released, dipeptidyl peptidase IV (DPP-IV) cleaves the N-terminal tyrosine-proline
residues to form PYY3-36 [186]. In humans, PYY is expressed in L cells in the lower part of the
ileum, colon, and rectum [187]. Similar to GLP-1, PYY is released in two phases, with the early
phase occurring when the nutrients are in the proximal gut [188], and have not reached the ileum
or colon yet. Indeed, PYY can be stimulated by the infusion of fat into the duodenum in humans
[32;33;58;189]. In response to fat, fat hydrolysis [32;33] must occur to release LCFAs
[34;60;190] for PYY secretion to be stimulated. Given the short timeframe for the first phase of
PYY release, a neural mechanism connecting the proximal gut and the distal gut is highly likely
to be in place. CCK has been suggested to mediate the release of GLP-1 stimulated by duodenal
nutrients, and there is evidence to suggest that CCK-A receptor signaling may be responsible for
the release of PYY stimulated by duodenal lipids [32].
In the circulation, both forms of PYY are present, but they can bind to different sets of receptors.
PYY1-36 binds to the Y1, Y2, Y4 and Y5 receptors, and PYY3-36 predominantly binds to the Y2 and
Y5 receptor [191]. Since DPP-IV is very widespread in the body, PYY3-36 is the dominant form
in the circulation, and PYY3-36 seems to play an important role in the regulation of glucose
homeostasis. In contrast to the other gastrointestinal peptides that are released postprandially to
lower glucose levels, PYY3-36 infusion increases postprandial glucose concentrations [192]. The
rather different effect of PYY3-36 on postprandial glucose levels from the other gastrointestinal
peptides may be attributed to its inhibitory effect on GSIS [193], preventing the secretion of
insulin to lower glucose. Moreover, PYY3-36 has no effects on insulin sensitivity under basal
21
conditions [194]. Hence, PYY3-36 may be responsible for balancing the effects of other
gastrointestinal peptides to prevent hypoglycemia as a result of the augmented postprandial
insulin secretions.
In addition to increasing postprandial glucose levels, both PYY1-36 and PYY3-36 can inhibit
gastric emptying with PYY3-36 being more effective [195;196]. These findings suggest that
PYY3-36 can also exert suppressive effects on glucose levels secondary to gastric emptying.
Similar to GLP-1, PYY3-36 has been shown to reduce food intake in rats [174]. In response to
PYY3-36, the arcuate nucleus of the hypothalamus is activated and both the food intake and brain
activation effects are abolished in rats subjected to subdiaphragmatic vagotomy or transection of
a brainstem-hypothalamus pathway [174]. These findings suggest that PYY3-36 can regulate
energy homeostasis through a neuronal network, supporting the existence of a gut-brain neuronal
axis.
22
1.4 Neuronal Regulation of Glucose Homeostasis by the Small Intestine
As mentioned, the small intestine can regulate glucose homeostasis through the actions of
gastrointestinal peptides as well as the activation of neuronal networks by nutrients. While it
appears that the mechanisms of action for gastrointestinal peptides are largely endocrine in
nature, some gastrointestinal peptides can also activate neuronal networks. For example, portal
GLP-1 can stimulate a hepatopancreatic reflex pathway to stimulate insulin secretion [169].
Similarly, portal administration of glucose can also stimulate hepatic glucose uptake and
peripheral glucose utilization [18] via neural communications.
Moreover, while some gastrointestinal peptides can regulate energy homeostasis through a gut-
brain neuronal axis, this axis is also involved in the inhibition of food intake by upper intestinal
lipids [197]. Recently, our laboratory has discovered a gut-brain-liver axis induced by duodenal
lipids to suppress glucose production to regulate glucose homeostasis [20].
In the following section, the neuronal axes involved in the regulation of glucose homeostasis
activated in the gut will be discussed.
23
1.4.1 Hepatoportal Glucose Sensor
As glucose is absorbed from the intestine, glucose can enter the portal vein. It has been suggested
that as glucose enters the portal vein, the body can adopt appropriate mechanisms to compensate
for the incoming glucose. Indeed, a hepatoportal glucose sensor has been identified to stimulate
hepatic glucose uptake and peripheral glucose utilization in independent studies.
Hepatic glucose uptake
Studies on the effect of portal glucose on hepatic glucose uptake were mainly carried out by the
group of Cherrington AD. In brief, they discovered that hepatic glucose uptake is increased when
glucose is delivered intraportally in comparison to peripheral vein administration in dogs [198].
Importantly, the increase in hepatic glucose uptake was blocked when the liver was denervated
[199], suggesting that neural communications is required for the effect. The neural component of
this effect was proposed to be of parasympathetic nature based on two observations. First, it was
found that afferent discharges in the hepatic branch of the vagus nerve can be modulated by
portal glucose in guinea pigs [200]. Second, acetylcholine administration in the portal vein was
also able to stimulate hepatic glucose uptake in dogs [201]. In the liver, the glucose taken up is
disposed as glycogen in dogs [202] and rats [203]. Hence, a vagally-mediated mechanism exists
to mediate communication between the portal vein and the liver to upregulate hepatic glucose
uptake in response to portal signals.
24
Peripheral glucose utilization
As mentioned, a hepatoportal glucose sensor exists to stimulate peripheral (muscle) glucose
utilization in response to portal glucose infusion in mice [18]. Since the hepatoportal glucose
sensor is connected to the hypothalamus and the nucleus of the solitary tract through hepatic
vagal afferents [204-207], a vagus-mediated neuronal network is likely in place between the
portal vein and the brain. Moreover, the effect of intraportal glucose infusion was abolished in
denervated muscles, suggesting that neuronal communication between the brain and oxidative
tissues is also necessary to increase glucose utilization [18].
While these findings formulate a basis for the portal-brain-muscle neuronal axis, the mechanism
of activation at the level of the portal vein appears to require the mediation by GLP-1 since the
effect of this neuronal axis is absent in GLP-1 receptor knockout mice [208]. At the level of the
muscle, the increase in glucose utilization is attenuated in mice lacking the expression of glucose
transporter-4 (GLUT4) or mice expressing the dominant negative form of adenosine
monophosphate kinase (AMPK) [209], suggesting that the activation of AMPK and the presence
of GLUT4 are required to increase glucose utilization by the muscles. In contrast, muscle-
specific inactivation of the insulin receptor gene had no effects on the stimulation of glucose
utilization, implying that the effect operates in an insulin-independent manner [209].
Nonetheless, a glucose-induced portal-brain-muscle axis exists to regulate glucose homeostasis
through enhancing glucose utilization by oxidative muscles.
25
1.4.2 Lipid-induced gut-brain-liver axis
Recently, our laboratory has demonstrated that intraduodenal administration of lipids can trigger
a gut-brain-liver axis to suppress glucose production [20].
At the level of the duodenum, the lipid-sensing mechanism is dependent on lipid metabolism. As
lipids in the form of triglycerides enter the duodenum, the triglycerides are hydrolyzed by lipases
to release fatty acids. As fatty acids, specifically LCFAs, are released, the LCFAs are converted
into long-chain fatty acyl-coenzyme A (LCFA-CoA) by acyl-coenzyme A synthetase (ACS), and
the accumulation of LCFA-CoA in the duodenum is required to lower glucose production. This
is evident by the fact that when lipids are co-infused with the ACS inhibitor, triascin C, the
ability of upper intestinal lipids to lower glucose production was abolished. Hence, lipid-sensing
in the duodenum requires the esterification of lipid metabolites.
Further experiments demonstrated that a neuronal axis connecting the gut to the brain, then the
brain to the liver is involved. Briefly, duodenal lipids failed to lower glucose production in the
presence of tetracaine (a local anesthetic which inhibits neuronal activation), indicating that
duodenal lipids regulate glucose production in the preabsorptive state by activating
neurotransmissions. It was identified that the vagal afferents innervating the gut is responsible
for mediating the signals because intraduodenal lipid infusion in rats that received vagal
deafferentations did not lower glucose production. The critical role of the vagus nerve in
mediating the lipid-induced effect was further confirmed when glucose production remained
unchanged in response to intraduodenal administration of lipids in rats that received a
26
subdiaphragmatic vagotomy. A hindbrain region known as the nucleus of the solitary tract (NTS)
was identified to be the receiver of the vagal signals [210-213]. Importantly, the principal
neurotransmitter released from vagal afferent terminals in the NTS is glutamate [214] [215].
Glutamate can act on at least 4 different classes of receptors: NMDA, α-amino-3-hydroxy-5-
methyl-4-isoxazolproprionic acid (AMPA), kainite, and metabotropic receptors [216]. In the
NTS, NMDA receptors have been localized [217], and a previous study has demonstrated that
the administration of MK-801 (N-methyl D-aspartate (NMDA) receptor blocker) in the NTS
extended meal duration [218]. As such, MK-801 was administered in the NTS concomitantly
with the administration of lipids in the duodenum to assess whether NMDA receptors in the NTS
play a role in the lipid-induced glucose production suppressive effect. It was found that duodenal
lipids had no effects on glucose production, suggesting that the gut sends signals to the NTS
through the activation of NMDA-receptors in response to lipids. Lastly, intraduodenal
administration of lipids into rats that received a hepatic vagotomy failed to lower glucose
production, demonstrating the essentiality of the hepatic vagus nerve in mediating the effect.
Hence, after the NTS receives the signals from the gut, the signals are relayed to the liver to
lower glucose production via the hepatic vagus nerve. Ultimately, these experiments revealed a
gut-brain-liver axis activated by upper intestinal lipids to regulate glucose homeostasis (Figure
1). This neuronal axis represents one of the first lines of metabolic defenses against nutrient
excess to provide metabolic balance by lowering glucose production upon nutrient exposure.
Model of Diet-Induced Insulin Resistance
Since it is clear that a gut-brain-liver axis exists to regulate glucose production in response to
duodenal lipids in normal rodents, it was further evaluated whether the effect is present in a
27
model of diet-induced insulin resistance. It has previously been demonstrated that Sprague
Dawley (SD) rats can develop hepatic insulin resistance in response to three days of high fat
feeding [219]. In contrast to rodents fed on standard chow, intraduodenal administration of lipids
failed to lower glucose production in this model of diet-induced insulin resistance. Therefore, it
is essential to investigate the downstream mechanisms involved in mediating the activation of
the gut-brain-liver axis by duodenal lipids to uncover possible ways to restore the functionality
of this axis.
28
Figure 1 Upper intestinal lipids activate a gut-brain-liver axis to suppress glucose production. As lipids enter the duodenum (first segment of the small intestine), lipids in the form of triglycerides are broken down by lipases to release long-chain fatty acids in the lumen. Long chain fatty acids (LCFA) are converted to long-chain fatty acyl-coenzyme A (LCFA-CoA) and the accumulation of LCFA-CoA leads to the activation of vagal afferents innervating the duodenum. The vagus nerve sends the signals to the nucleus of the solitary tract (NTS) through activating the N-methyl-D-aspartate (NMDA) receptors. The NTS then acts as a relay center and send signals through the hepatic vagus nerve to suppress glucose production.
lumen
Lamina propria
Vagus
DUODENUM lipid
Lipase
LCFA
ACS
CoA LCFA-CoA
NTS NMDA receptors
Glucose Production
Vagus
29
2 General Hypothesis and Aims Cholecystokinin
Here, we identify CCK as a possible candidate for the downstream signaling events underlying
the duodenal lipid-sensing process that leads to the suppression of glucose production via a gut-
brain-liver axis. While lipids are known to inhibit food intake, there is now accumulating
evidences suggesting that lipid-induced CCK mediates the satiety effect through a neuronal
network in the preabsorptive state, suggesting that CCK can activate vagal afferents to send
signals to the brain [220;221]. In food intake studies, it has been shown that the activation of
duodenal CCK-A receptors is responsible for mediating the lipid-induced satiation.
Pharmacological blockade of CCK-A receptors by its specific antagonist MK-329 has been
shown to reduce lipid-induced satiation [222-225]. Additionally, duodenal lipids fail to lower
food intake in CCK-A receptor deficient (OLETF) rats [226;227]. Together, studies utilizing
either pharmacological or genetic means to prevent the activation of CCK-A receptors in the
presence of duodenal lipids demonstrated that CCK and CCK-A receptors play a significant role
in mediating duodenal lipid-sensing mechanisms.
Duodenal CCK-induced gut-brain-liver axis
The general hypothesis of this thesis is that duodenal CCK can activate a gut-brain-liver axis to
regulate glucose homeostasis (Figure 2). First it will be assessed whether CCK-A receptors play
a role in the regulation of glucose production. Secondly, duodenal lipids have been shown to
activate a gut-brain-liver axis to suppress glucose production while the duodenal lipid-sensing
30
mechanisms remain unknown. Since CCK-A receptors have been located on capsaicin-sensitive
vagal afferents [228;229] and the duodenal lipid-induced glucose production lowering effect is
dependent on vagal afferent signaling, it is plausible that duodenal CCK/CCK-A receptor
signaling is downstream of lipid-sensing. In order for us to be able to examine whether duodenal
CCK can regulate glucose homeostasis independent of changes in glucose-regulatory hormones,
we will be utilizing clamp studies in which we will fix insulin levels at near-basal levels.
OLETF – CCK-A receptor deficient rats
One model that is available to evaluate whether CCK-A receptors are involved in the regulation
of glucose homeostasis is a rat strain known as Otsuka Long Evans Tokushima Fatty (OLETF)
rats. OLETF rats are an outbred strain of Long-Evans rats that congenitally lack a 6847 base pair
segment of the promoter region of the gene encoding CCK-A receptors [230]. Consequently,
OLETF rats are deficient in CCK-A receptors. Importantly, CCK-A receptors are involved in the
regulation of feeding behaviour, thus OLETF rats are hyperphagic from birth. When placed on a
regular chow ad libitum, OLETF rats develop glucose intolerance by week 5, obesity by week
10, and diabetes by week 24 [231-233] as compared to the LETO control rats. Of relevance to
this study, it has been shown that at week 8, OLETF rats have a higher basal hepatic glucose
production than LETO rats when fed ad libitum [234]. This finding supports our hypothesis that
duodenal CCK signaling is necessary to negatively regulate glucose production.
Since OLETF rats tend to be obese by week 10, in order to evaluate whether duodenal CCK
regulates glucose production in OLETF rats independent of weight gain, we will pair-feed the
31
OLETF rats to LETO rats once we receive them from Tokushima Institute, Otsuka
Pharmaceutical at 4 weeks of age. Subsequently, we will be able to evaluate whether duodenal
CCK-A receptor plays a role in the regulation of glucose production by using OLETF rats that
are pair-fed with LETO rats.
Physiological relevance
After confirming the role of duodenal CCK on the regulation of glucose homeostasis using
clamp studies, we will evaluate whether duodenal CCK regulates plasma glucose levels in
response to fasting-refeeding in rats to address the physiological relevance of our findings. Since
we are hypothesizing that CCK acts downstream of duodenal lipid-sensing to regulate glucose
homeostasis, feeding will provide a physiological means to introduce nutrients into the
duodenum. By performing a fasting/refeeding protocol, we will be able to assess whether
blocking duodenal CCK signaling will impair the regulation of plasma glucose levels in
physiological settings. Altogether, these experiments will allow us to determine whether
duodenal CCK can regulate glucose production in normal settings.
High-fat diet model
Moreover, we will also evaluate whether duodenal CCK can regulate glucose production in the
high-fat diet-induced model of early onset insulin resistance. Based on the findings from feeding
studies, upper intestinal lipids fail to regulate energy homeostasis following adaptation to high
fat feeding [235], consistent with our previous findings in regards to glucose homeostasis. It has
32
been proposed that the dysregulation of energy homeostasis by upper intestinal lipids is due to a
defect in CCK signaling. In both rats and humans, high-fat diets have been associated with
elevated levels of circulating CCK [236;237]. This observation leads to the speculation that
individuals who consume high-fat diets chronically may develop CCK insensitivity, such that an
upregulated level of CCK is required to elicit satiation in response to upper intestinal lipids. This
hypothesis is further supported by the finding that exogenous CCK reduced food intake
significantly less in rats fed with high-fat diets for 3 weeks compared with rats fed with low-fat
diets [238-240]. Hence, these findings suggest that CCK insensitivity develop following the
adaptation to high-fat feeding. In regards to the regulation of glucose homeostasis, we
hypothesize that CCK-resistance will impair the regulation of glucose homeostasis by duodenal
CCK in response to high-fat feeding.
33
Hypothesis: Duodenal CCK can suppress glucose production through the activation of a gut-
brain-liver axis.
Aim 1: To determine whether the activation of CCK-A receptors in the duodenum can suppress
glucose production.
Aim 2: To examine whether duodenal CCK can stimulate a gut-brain-liver axis to suppress
glucose production.
Aim 3: To investigate whether CCK/CCK-A receptor signaling is required for duodenal lipids
to lower glucose production.
Aim 4: To confirm whether the regulation of glucose homeostasis by duodenal CCK is
physiologically relevant.
Aim 5: To determine whether duodenal CCK can lower glucose production in a model of diet-
induced insulin resistance.
34
Figure 2 Working hypothesis – upper intestinal lipids stimulate CCK/CCK-A receptor signaling to suppress glucose production through a gut-brain-liver axis As lipids in the form of triglycerides enter the lumen of the duodenum as part of the ingested food, CCK is released and it binds to CCK-A receptors (CCK-AR) on the vagus nerve to send signals to the NTS by activating the NMDA receptors. The NTS acts as a relay center and send signals through the hepatic vagus nerve back to the periphery to suppress glucose production.
Vagus
lumen
Lamina propria
DUODENUM lipid
Lipase
LCFA
ACS
CoA LCFA-CoA
NTS NMDA receptors
Glucose Production
Vagus
CCK-AR CCK
35
3 General Materials and Methods
*Note: All the animal study protocols were reviewed and approved by the Institutional Animal
Care and Use Committee of the University Health Network.
Chemicals
CCK-8 (Sulfated), tetracaine, and NMDA receptor blocker MK-801 were obtained from Sigma.
CCK-A receptor antagonist MK-329 was obtained from Tocris Bioscience. 20% Intralipid was
obtained from Baxter Healthcare Corporation. Stock solution of CCK-8 and MK-801 were
prepared in saline whereas stock solutions of tetracaine and MK-329 were prepared in
dimethylsulfoxide (DMSO). All stock solutions were diluted in 0.9% NaCl to the desired
concentrations.
Models
Male Sprague-Dawley Rats
9-week old Sprague-Dawley (SD) rats, weighing between 250 – 300g (Charles River
Laboratories, Montreal QC) were used for our studies. Rats were housed individually and
maintained on a standard 12-12h light-dark cycle with access to rat chow (Harlan Teklad 6%
Mouse/Rat Diet; composition: 52% carbohydrate, 31% protein and 17% fat; total calories
provided by digestible nutrients: 3.83 kcal/g) and water ad libitum. The rats were allowed to
acclimatize for 5 days upon arrival and the appropriate surgeries were then performed.
36
Male LETO/OLETF Rats
4-week old Long Evans Tokushima Otsuka (LETO) and Otsuka Long Evans Tokushima Fatty
(OLETF) rats were obtained as a generous gift from Dr. Kawano (Tokushima Research Institute,
Otsuka Pharmaceuticals, Tokushima, Japan). Rats were housed individually and maintained on a
standard 12-12h light-dark cycle with water ad libitum. In order to avoid the results from being
confounded with the effects of obesity, OLETF rats were pair-fed with LETO rats to prevent the
development of obesity as described [241]. All LETO rats had access to standard chow (Harlan
Teklad 6% Mouse/Rat Diet as described) ad libitum. At 8AM each day, body weights and food
intakes of both LETO and OLETF rats were measured to ensure that the body weights were
similar. After daily food intakes of LETO rats were measured, OLETF rats were supplied with
the amount of chow equal to the prior day’s average daily chow intake of LETO rats maintained
on ad libitum access. The OLETF rats were pair-fed until they were 11-12 weeks old (280-300g),
and the appropriate surgeries were then performed.
Surgical Procedures
Duodenal and Intravenous Cannulations
Rats were first anesthetized with intraperitoneal (ip) ketamine (Ketalean; Bimeda-MTC,
Cambridge, Ontario) and xylazine (Rompun; Bayer). Duodenal cannulation surgeries were
performed as described [20;227]. Laparotomy incisions were made on the ventral midline and
the abdominal muscle wall to expose the gastrointestinal tract in the peritoneum. The pyloric
sphincter was identified and the proximal 1.5cm of the duodenum was then isolated. A 25-gauge
37
needle was used to make a small puncture would on the ventral aspect of the duodenum in a
region where the vascular arcade was as sparse as possible to minimize bleeding. A saline-filled
catheter made of silicone tubing (0.040 in. ID, 0.085 in. OD; Sil-Tec, Technical Products, USA)
with a 0.5cm extension of a smaller silicone tubing (0.025 in. ID, 0.037 in. OD; Sil-Tec,
Technical Products, USA) was inserted into the duodenum at a position approximately 2cm
downstream of the pyloric sphincter for intraduodenal infusions [20]. The cannula was flushed
with saline to ensure that it was inserted into the lumen of the duodenum. The cannula was
anchored to the outer serosal surface of the proximal duodenum around the puncture wound with
a drop of 3M adhesives (Vetbond) and a 0.5-cm2 piece of Marlex mash sewn to the serosal
surface with a 6-0 silk suture. The proximal portion of the cannula exited to the abdominal cavity
through the site of the laparotomic incision and the abdominal wall incision was closed with a 4-
0 silk suture. A 2-cm midline incision was made on the skin of the back of the neck, just rostral
to the interscapular area, and the proximal portion of the duodenal cannula was then tunneled
subcutaneously to exit through the incision. All the skin incisions were closed with 4-0 silk
sutures and the proximal end of the duodenal cannula was closed with a metal pin.
For rats that were to undergo the pancreatic (basal insulin) – euglycemic clamp experiments,
immediately following the duodenal cannulation surgery, indwelling catheters were also inserted
into the right internal jugular vein and left carotid artery for infusion and blood sampling
purposes. Catheters were made of polyethylene catheters (PE 50, Clay Adams) with a 15mm
cuff-extension of Silastic tubing (Corning). Both catheters were tunneled subcutaneously and
exteriorized. The catheters were filled with 10% heparinzed saline to maintain potency of the
vascular cannula, and then closed at the end with a metal pin.
38
Recovery from surgery was monitored by measuring daily food intake and weight gain for 4-6
days after surgery and the duodenal catheter was flushed with saline daily to ensure potency.
Stereotaxic Surgery
For a subgroup of rats which required stereotaxic surgeries, rats were stereotaxically implanted
with indwelling cannula (Plastics One Inc., Roanoke, VA) according to the atlas of the rat brain
as previously described [20]. In brief, rats were anesthetized with ip ketamine and xylazine then
fixed in a stereotaxis apparatus (David Kopf Instruments, Tujunga, CA) with ear bars and a nose
piece set at +5.0mm. 26-gauge stainless steel double guide cannulae were used for implantations
in the nucleus of the solitary tract with the following coordinates [20]: 0.0mm on occipital crest,
0.4mm lateral to midline, 7.9mm below skull surface. Rats were given a week of recovery time
post-stereotaxic surgery in individual cages, maintained on a standard 12h-12h light-dark cycle
with access to standard rat chow and water ad libitum, followed by duodenal and vascular
cannulation surgeries.
Selective Hepatic Branch Vagotomy
For a subgroup of rats which were subjected to hepatic branch vagotomy, the surgeries were
performed as previously described [20]. A laparotomy incision was made on the ventral midline,
followed by a second incision to open the abdominal muscle wall, exposing the gastrointestinal
tract in the peritoneum. A gastrohepatic ligament was severed using fine forceps and the stomach
was gently retracted into sterile saline soaked cotton gauze to reveal descending esophagus and
39
ventral subdiaphragmatic vagal trunk. The hepatic branch of the ventral subdiaphragmatic vagal
trunk was transected by microcautery in between the two sutures, severing and cauterizing
hepatic vagus. Transecting the hepatic branch of the vagal nerve disrupts neural communications
between the liver and the brain. This also results in slightly decreased innervations to the
intestine as there are minor innervations of the hepato-duodenal sub-branch which supplies a
small portion of the intestine. Following the vagotomy surgery, duodenal and vascular
cannulation surgeries were immediately performed.
High-Fat Feeding in Male SD Rats
A subgroup of male SD rats, with duodenal and vascular catheters implanted, was placed on a
lard-oil enriched high-fat diet ad libitum for 3 days (see Table 1 for diet composition of standard
chow and high-fat diet). It was first confirmed that the rats on high fat diet consumed at least the
same amount of calories as the average calories consumed for the rats on regular chow which
were used for other clamp experiments. A pancreatic basal insulin clamp was then performed on
these rats to address whether intestinal CCK regulates glucose production in the early onset of
diet-induced insulin resistance in rodents.
40
Intraduodenal Infusions and Treatments
The following substances were continuously infused to the duodenum through the duodenal
catheter at t = 150 – 200 min (0.01 mL/min):
1.) saline
2.) CCK-8 (35 pmol/kg/min; 100 pmol/kg/min)
3.) CCK-8 (35 pmol/kg/min) + tetracaine (0.01 mg/min)
4.) CCK-8 (35 pmol/kg/min) + CCK-A receptor antagonist MK-329 (1.6 µg/kg/min; 3.2
µg/kg/min)
5.) 20% Intralipids (0.03 kcal/min)
6.) 20% Intralipids (0.03 kcal/min) + MK-329 (1.6 µg/kg/min).
Since this is one of the first studies to examine the biological function of duodenal CCK-8 in
rats, we used an infusion rate that is analogous to the ip dose of CCK-8 at 1750 pmol/kg which
was found to suppress food intake in SD rats [242]. It was determined that CCK-8 was to be
infused at a rate of 35 pmol/kg/min (or 1750 pmol/kg in 50 min) as this rate is analogous to the
ip dose of CCK-8 at 1750 pmol/kg which was found to suppress food intake in SD rats [55]. This
infusion was further justified by a previous study which indicated that intraduodenal CCK-8
administered at 30 or 100 pmol/kg/min stimulates pancreatic secretion in calves independent of
changes in circulating CCK levels [243].
41
Fasting-Refeeding Experiments
For male SD rats, five days post-duodenal surgery, rats whose daily food intake and body weight
had recovered back to baseline underwent the fasting-refeeding protocol. Rats were fasted for 40
hours (fast began at 5:00 PM on Day 5 until 9:00 AM on Day 7). Ten minutes prior to the
completion of the 40-hour fast (i.e. 8:30 AM on Day 7, t = -10), a continuous intraduodenal
infusion (Harvard Apparatus PHD 2000 infusion pumps) of either (i) saline or (ii) MK-329 (3.2
µg/kg/min) was initiated, which lasted until t = 20. Upon completion of the 40-hour fast (i.e.
9:00 AM on Day 7, t = 0), rats were allowed to consume a regular chow diet ad libitum. Blood
glucose levels and food intake were measured at t = -10, 0, 10, 20 min.
On the other hand, male LETO and OLETF rats did not receive any surgeries. Similar to the
male SD rats, they were fasted for 40 hours. Upon completion of the 40-hour fast, LETO and
OLETF rats were allowed to consume a regular chow diet ad libitum. Blood glucose levels and
food intake were measured at t = 0, 10, 20 min.
Clamp Procedure
Rats were restricted to ~58 kcal of caloric intake the night prior to the in vivo infusion
experiments. The infusion experiments spanned a total of 200 minutes. At the onset of the
experiment, a primed continuous infusion of [3-3H]-glucose (bolus 40 µCi; 0.4 µCi/min; all
infusion performed with Harvard Apparatus PHD 2000 infusion pumps) was initiated (t = 0 min)
and maintained throughout the protocol to assess glucose kinetics based on the tracer-dilution
methodology.
42
At t = 90 min, a pancreatic (basal insulin) clamp was initiated by providing a continuous infusion
of insulin (0.8 mU/kg/min) and somatostatin (3 µg/kg/min) to inhibit endogenous insulin and
glucagon secretions. To maintain euglycemia, 25% glucose was provided at a variable rate to
maintain plasma glucose levels that are comparable to basal (t = 60 – 90 min), and the rates were
adjusted at 10-min intervals from t = 120 – 200 min. For experiments which required the
administration of MK-801 (0.03 ng/min; CMA/400 syringe microdialysis infusion pumps) to the
NTS, the infusion began at t = 90 min and lasted until t = 200 min. Intraduodenal infusions were
initiated at t = 150 min and lasted until t = 200 min to determine the effects of different duodenal
treatments on glucose kinetics. Blood samples were taken at 10-min intervals to determine the
specific activity of [3-3H]-glucose between t = 60 – 90 min (basal) and t = 180 – 200 min
(clamp) to assess glucose kinetics. Additional blood samples were collected at t = 90 min (basal)
and t = 180 and 200 min to determine plasma insulin levels. All the blood samples were
subjected to centrifugation at 6000 rpm to separate the plasma and the biochemical analyses were
performed as described below.
Peripheral and Portal CCK Measurements
A commercially available radioimmunoassay kit (Alpco) was used for the measurement of CCK
concentrations in the peripheral and portal circulation. According to the manufacturer’s
specifications, 1.0 mL of plasma was required to obtain a precise CCK measurement. Therefore,
a separate set of experiments was required to collect sufficient plasma for the measurement of
CCK. In this set of experiments, SD rats underwent both the duodenal and vascular cannulation
surgeries. Following a recovery period, they received either saline or CCK-8 in the duodenum to
43
determine whether duodenal saline or CCK-8 has any effect on peripheral and portal CCK
concentrations. The infusion lasted for 50 minutes to mimic the protocol used in the clamp
studies. At t = 50 min, 2.0 mL of blood was collected. The rats were anesthetized immediately
using ketamine (5 mg IV) and 2.0 mL of portal blood was extracted from the portal vein. The
blood samples were put in tubes containing 1 unit of ethylenediaminetetraacetic acid (EDTA;
(Phoenix Pharmaceuticals) and 1 unit of aprotinin (Phoenix Pharmaceuticals) and plasma was
separated by centrifugation at 6000 rpm for 5 minutes at 4ºC. Plasma samples were stored in -
20ºC until the radioimmunoassay was performed.
Biochemical Analyses
Plasma Glucose
Plasma glucose concentrations were measured with the use of a glucose analyzer (Glucose
Analyzer GM9, Analox Instruments, Lunenbertg, MA). The analyzer was calibrated before
usage in each experiment. Blood samples of rat (~0.1mL) were centrifuged at 6000 rpm to
separate the plasma. A 10 µL sample of plasma was immediately injected into the glucose
analyzer to determine plasma glucose concentration. The glucose analyzer determines glucose
concentrations by the glucose oxidase method, which is based on the following reaction:
Under the assay conditions, the rate of oxygen consumption is directly proportional to the plasma
glucose concentration. Therefore, oxygen consumption is measured in the glucose analyzer with
a polarographic oxygen sensor to determine the plasma glucose concentration. Specifically,
Clark-type amperometric oxygen electrodes are immersed in the sample with a potential applied
β-D-glucose + O2 D-Gluconic acid + H2O2 Glucose oxidase
44
between them that is sufficient to reduce dissolved oxygen at the working electrode. Through
this, the partial pressure of oxygen in the sample can be measured given that it is proportional to
the limiting current.
Plasma Glucose Tracer Specific Activity
The specific activity of the [3-3H] in plasma was determined using 50-µL samples of plasma.
The plasma samples were deproteinized with Ba(OH)2 and ZnSO4, and centrifuged at 6000 rpm
for 7 minutes at 4ºC. The protein-free supernatant was kept. Since tritium on the C-3 position of
glucose is lost to water during glycolysis, the supernatant was evaporated to dryness to remove
the tritiated water. Scintillation fluid was added to the dried sample and the liquid scintillation
counting would represent radioactivity from the [3-3H]-glucose in the plasma only.
Plasma Insulin
Plasma insulin levels were determined by radioimmunoassay (RIA) using a 2-days commercial
rat insulin RIA kit (100% specificity) from Linco Research (St. Charles, MO). The principle of
RIA is based on antigen-antibody binding. In brief, insulin from the plasma samples or standards
competes with the labeled tracer antigen (125I-labeled insulin) to bind with the antibodies raised
against insulin (guinea pig anti-rat insulin antibody). As the reaction occurs, 125I-labeled insulin
binds in a reverse proportion to the concentration of insulin in standards and samples. Antibody-
bound labeled tracer antigens (125I-labeled insulin) are separated from the unbound fraction using
double antibody solid phase. The radioactivity of the bound fraction is measured in a gamma
45
counter. The radioactivity counts (B) for the samples and the standards are expressed as a
percentage of the mean counts of the total binding reference tubes (B0):
The percent activity bound for each standard is plotted against the known concentration to
construct a standard curve. Finally, the concentration of insulin in the samples can be determined
by interpolation.
Specifically, a 2-days protocol provided by the supplier was used. 125I-labeled insulin (50 µL)
and rat insulin antibody (50 µL) were added to 50 µL of experimental samples and standards in a
range of concentrations (0.1, 0.2, 0.5, 1.0, 2.0, 5.0 and 10.0 ng/mL), followed by vortexing. After
an overnight incubation at 4ºC, 1.0mL of precipitating reagent was added to each tube followed
by vortexing and incubation at 4ºC for 20 minutes. The samples were centrifuged to pellet the
bound insulin. The radioactivity of the pellet was counted by a gamma counter (Perkin Elmer
1470). A standard curve was constructed using the method as described previously and the
concentration of insulin in the samples was determined by interpolation.
Plasma CCK
Peripheral and portal CCK levels were measured using a commercially available
radioimmunoassay kit (Alpco). This RIA kit follows the same principle as described in the
plasma insulin section. Briefly, CCK from the peripheral and portal plasma samples or standards
competes with the labeled tracer antigen (125I-labeled CCK) to bind with the antibodies raised
against CCK (rabbit anti-CCK antibody). As the reaction occurs, 125I-labeled CCK binds in a
% total binding = % = x 100% B0 B
Sample or standard B0
46
reverse proportion to the concentration of CCK in standards and samples. Antibody-bound
labeled tracer antigens (125I-labeled CCK) are separated from the unbound fraction using double
antibody solid phase. The radioactivity of the bound fraction is measured in a gamma counter.
The radioactivity counts (B) for the samples and the standards are expressed as a percentage of
the mean counts of the total binding reference tubes (B0). A standard curve is constructed by
plotting the percent activity bound for each standard against the known concentration. Finally,
the concentration of CCK in the samples is determined by interpolation.
Prior to the performance of the RIA, CCK was first extracted from the plasma samples to
eliminate non-specific interference from plasma proteins. 2.00 mL of 96% ethanol was added to
the plasma samples, vortexed. The samples were allowed to incubate for 10 minutes, followed by
centrifugation at 1,700g for 15 minutes. The CCK-containing supernatant was kept and allowed
to dry by using a Speed Vac (Savant SPD 131 DDA) overnight. The dry extracts were dissolved
back into solution by adding 1.00 mL of diluent. The dissolved samples were vortexed and
incubated for 30 minutes in room temperature. A recovery control for the extraction procedure
was also included by adding 200 µL of CCK-standard (50 pmol/L) to 800 µL to separate plasma
samples, making the final concentration 10 pmol/L. 200 µL of diluent was added to another 800
µL of plasma sample and both of these samples underwent the same extraction procedure and
RIA with the experimental samples concurrently. The inclusion of the recovery controls allowed
for the determination of the recovery rate of the extraction procedure, which could be calculated
by the following formula:
47
After calculating the % recovery from the recovery controls, the % recovery was used to correct
the concentration of CCK in the samples as determined by the RIA.
For the RIA, a 7-days protocol provided by the supplier was used. 500 µL of 125I-labeled CCK
and 500 µL of anti-CCK were added to 200 µL experimental samples and standards in a range of
concentrations (0.78, 1.56, 3.12, 6.25, 12.5, 25 pmol/L). The tubes were vortexed and allowed to
incubate for 4 days at 4ºC. 100 µL of precipitating reagent was added to each tube followed by
vortexing and incubation at 4ºC for an hour. The samples were centrifuged to pellet the bound
CCK. The radioactivity of the pellet was counted by a gamma counter (Perkin Elmer 1470). A
standard curve was constructed and the final concentration of CCK in the samples was
determined by interpolation and accounting for the % recovery.
Calculations
As described previously, a radioactive [3-3H]-glucose tracer is infused during the clamp
experiments in order to determine glucose production and uptake in our experimental animals
using the steady state formula. The [3-3H]-glucose tracer was infused at a constant rate into the
rat and an hour was given to allow for equilibration of the tracer glucose with the glucose in the
body. Under steady-state basal condition, the rate of glucose uptake (Rd) equals the rate of
glucose appearance (Ra), which is the same as the rate of the endogenous glucose production.
Therefore, using the steady state formula, the rate of glucose uptake and glucose appearance can
% recovery = x 100% pmol/L found with addition – pmol/L found without addition
10 pmol/L
48
be determined by dividing the [3-3H]-glucose infusion rate by the specific activity of the plasma
[3-3H]-glucose:
During the pancreatic clamp settings where exogenous glucose was infused to maintain
euglycemia, the rate of endogenous glucose production was obtained from the difference
between Rd and the rate of glucose infusion.
Statistical Analysis
Data were presented as means ± standard errors of the mean. Statistical differences between
groups were determined by either analysis of variance (ANOVA) followed by Tukey’s test or the
unpaired Student’s t-test as appropriate with a probability of p < 0.05 accepted as significant. For
pancreatic clamp experiments, the time period t = 60 – 90 min and t = 180 – 200 min were
averaged for the basal and clamp condition respectively.
Ra = Rd = Constant tracer infusion rate (µCi/min)
Specific activity (µCi/mg)
49
Calories Provided Regular Chow Diet High-Fat Diet
Carbohydrate (%) 52 45
Protein (%) 31 22
Fat (%) 17 33
Saturated 6.2 14.1
Monounsaturated 7.1 13.2
Polyunsaturated 3.5 6.1
Total calorie provided (kcal/g) 3.83 5.14
Table 1 Dietary contents of the regular chow and the lard-oil enriched high fat diet.
50
4 Results
Duodenal CCK-A receptor activation can suppress glucose
production
During the clamp studies (180 – 200 min), when peripheral circulating plasma insulin was
maintained at near basal levels, intraduodenal CCK-8 administration (Figure 3A, B) significantly
increased the exogenous glucose infusion rate required to maintain euglycemia in comparison to
saline (p < 0.05) (Figure 4A). Based on the steady-state tracer data (180-200 min), this was
selectively attributed to a significant reduction in glucose production (Figure 4B,C) in rats that
received CCK-8 (p < 0.001) with no changes in glucose uptake (Figure 4D). In another set of
rats, we performed intraduodenal CCK-8 (35 pmol/kg/min) administration for 50 min and found
plasma CCK levels did not change at 50 min. Portal vein samples were then taken soon after
giving anesthesia at 50 min. It was found that CCK levels in the portal vein (5.1 ± 0.6 pM) was
~2.2-fold higher than the plasma CCK levels obtained in duodenal saline-infused rodents,
without reaching statistical significance. Importantly, portal CCK levels of duodenal CCK-8
infused rats (3.4 ± 0.6 pM) were comparable to those found in the saline-infused rats. Thus, a
primary increase of CCK-8 in the duodenum lowers glucose production independent of changes
in circulating CCK or insulin levels (Table 2).
Next, we co-administered the CCK-A receptor inhibitor MK-329 (3.2 μg/kg/min) with CCK-8
into the duodenum (Figure 3A) to inhibit CCK-A receptors in the gut in the presence of CCK
activation. In the presence of MK-329, the effects of duodenal CCK-8 administration on glucose
51
infusion rate and glucose production were abolished, with no effects on glucose uptake.
Importantly, MK-329 (a competitive inhibitor for CCK-A receptors) alone also had no effects on
glucose infusion rate, glucose production, and glucose uptake when given alone (Figure 4A-D).
The abolishment of the glucose production suppression effect of duodenal CCK in the presence
of the CCK-A receptor inhibitor suggests that CCK-A receptor activation is required for the
potent control of gut CCK-8 on glucose production.
Duodenal CCK fails to lower glucose production in CCK-A receptor
deficient rats
After identifying that CCK-A receptor activation can lower glucose production via a
pharmacological approach, we wished to confirm this finding in OLETF rats, which are rats with
congenital CCK-A receptor deficiency (Figure 5A). Since these rats are known to be
hyperphagic, actions were taken in order to evaluate whether duodenal CCK-8 regulate glucose
production in OLETF rats independent of weight gain. Upon the arrival of the OLETF rats from
Tokushima Institute, Otsuka Pharmaceutical at 4 weeks of age, we began pair-feeding the
OLETF rats with the LETO rats. As shown in Figure 6A, we successfully maintained similar
body weights between LETO and OLETF rats by pair-feeding (Figure 6B). After ensuring that
the OLETF rats were not obese in comparison to LETO rats, we performed the same set of
experiments in the OLETF and LETO rats as in SD rats.
Initially, we administered CCK-8 to the duodenum of LETO rats at 35 pmol/kg/min (same rate
as in SD rats; Figure 5A,B). However, there were no changes in glucose infusion rate, glucose
52
production and glucose uptake. Therefore, we doubled the dosage of CCK-8 to 70 pmol/kg/min,
and there were still no effects on glucose infusion rate, glucose production and glucose uptake.
Finally, we administered CCK-8 to the duodenum of LETO rats at 100 pmol/kg/min, and an
increased glucose infusion rate was required to maintain euglycemia as compared to
intraduodenal saline-treated LETO rats (p < 0.01) (Figure 6C). This was accounted for by a
reduction in glucose production (p < 0.05) (Figure 6D,E), with no changes in glucose uptake
(Figure 6F). In contrast, intraduodenal CCK-8 administrations in OLETF rats had no effects on
the glucose infusion rate required to maintain euglycemia (Figure 6C). There were also no
changes in glucose production (Figure 6D,E) and glucose uptake (Figure 6F) in response to
intraduodenal CCK-8 administrations in OLETF rats.
It is unclear why LETO rats required a higher dosage of CCK-8 to have an effect on glucose
production than SD rats. However, a previous study demonstrated that in the duodenum, the
level of CCK-A receptor gene expression is significantly higher in SD rats than LETO rats [244]
indicating that LETO rats may have a lower protein expression of CCK-A receptors in the
duodenum. Hence, a possible reason for the increased dosage of CCK-8 required for LETO rats
to lower glucose production as compared to SD rats is that there may be potentially reduced
availability of CCK-A receptors, suggesting that LETO rats may be less sensitive to CCK-8 in
the duodenum than SD rats.
Nonetheless, both the pharmacological loss-of-function in SD rats and genetic loss-of-function
experiments in CCK-A receptor-deficient rats indicate that CCK-A receptor activation is
sufficient for CCK action in the duodenum to lower glucose production.
53
Duodenal CCK can activate a gut-brain-liver axis to regulate
glucose production
Subsequently, we examined whether a gut-brain-liver axis is involved in duodenal CCK
signaling. First, we co-infused CCK-8 with tetracaine to determine whether duodenal innervation
is required for duodenal CCK to lower glucose production (Figure 7A,B). As shown in Figure
8A-D, duodenal CCK-8 administration had no effects on glucose infusion rate, glucose
production and glucose uptake in the presence of tetracaine. Importantly, the administration of
tetracaine alone had no effects on all parameters. Taken together, these results confirmed our
prediction that duodenal CCK regulates glucose production in the preabsorptive state by
activating neurotransmissions.
Next, we examined whether NMDA receptor activation in the NTS is necessary for duodenal
CCK to lower glucose production. We found that the effects of duodenal CCK-8 administration
on glucose infusion rate and glucose production were completely abolished when NMDA
receptor activation in the NTS is prevented by the administration of MK-801 (Figure 8A-C).
Consistently, glucose uptake was unchanged with concomitant administrations of CCK-8 in the
duodenum and MK-801 in the NTS (Figure 8D). MK-801 administration in the NTS alone had
no effects on all of the parameters. Hence, these data suggest that NMDA receptors in the NTS
play a role in mediating the glucose production suppression effect elicited by duodenal CCK.
After confirming that signals are transmitted from the gut to the brain, we examined whether the
hepatic vagus nerve plays a role in transmitting signals from the brain to the liver to lower
glucose production. We observed that the effects of duodenal CCK-8 administration on glucose
54
infusion rate and glucose production were abolished in rats that received a hepatic vagotomy
(Figure 8A-C), with no changes in glucose uptake in comparison to other groups (Figure 8D).
Notably, hepatic vagotomy had no effects on all of the parameters (Figure 8A-D). Together, our
results indicate that duodenal CCK activates a gut-brain-liver axis to regulate glucose
production.
Duodenal CCK-A receptor activation is required for lipids to trigger
a gut-brain-liver axis to regulate glucose production
Since CCK-A receptors have been implicated to play a role in mediating the lipid-induced gut-
brain axis to regulate energy homeostasis, we proposed that CCK-A receptors might also play a
role in mediating the lipid-induced gut-brain-liver axis to regulate glucose homeostasis (Figure
9A,B). First, we confirmed that duodenal lipids upregulated the glucose infusion rate required to
maintain euglycemia (p < 0.01) (Figure 10A), which is fully accounted by a suppression in
glucose production (p < 0.01) (Figure 10B,C) with no effects on glucose uptake (Figure 10D)
within the same timeframe as seen in CCK-infused experiments. We then co-infused lipids with
MK-329 into the duodenum to determine whether CCK-A receptor activation is required for
lipids to lower glucose production. In the presence of the CCK-A receptor inhibitor, duodenal
lipids had no effects on glucose infusion rate, glucose production and glucose uptake (Figure
10A-D). Hence, these data support the hypothesis that duodenal CCK-A receptor activation is
essential in mediating the lipid-induced gut-brain-liver axis to suppress glucose production.
55
Duodenal CCK-A receptors regulate plasma glucose levels in
physiological settings
To this point, we have demonstrated that duodenal lipid sensing can result in a reduction of
glucose production through the release of CCK which would activate a gut-brain-liver axis
through binding to duodenal CCK-A receptors. It is now important to evaluate whether these
findings are physiologically relevant by determining if CCK-A receptors are involved in the
regulation of plasma glucose levels in response to nutrient consumption.
To address this question, we performed a fasting-refeeding protocol, which allowed nutrients to
be delivered to the duodenum through a physiological mechanism to stimulate CCK release. Five
days after duodenal cannulation surgeries, the rats were subjected to a 40-hour fast (Figure 11A).
Following the fasting, we began an intraduodenal infusion of either saline or MK-329 at t = -10
min, during which the plasma glucose levels were at ~110 mg/dL (Figure 11B). At time 0 min,
we allowed the rats to feed ad libitum on regular chow, and plasma glucose levels remained the
same (Figure 11B). In response to feeding, both plasma glucose level and cumulative food intake
(Figure 11B,C) increased in 10 and 20 min of feeding for the intraduodenal saline-infused rats.
Interestingly, the plasma glucose levels in rats that received intraduodenal CCK-A receptor
blocker MK-329 were significantly higher than saline-treated rats by ~20 mg/dL at t = 20 min (p
< 0.01) (Figure 11B) despite a similar cumulative food intake (Figure 11C). Therefore, the
results of these pharmacological experiments suggest that duodenal CCK-A receptors in the
duodenum are involved in the regulation of glucose homeostasis in physiological settings.
56
Furthermore, we performed the same set of fasting-refeeding experiments in the CCK-A receptor
deficient OLETF rats and their corresponding LETO controls (Figure 12A). Similar to the
previous set of experiments, OLETF rats had significantly higher plasma glucose levels (Figure
12B) in comparison to LETO rats in response to 20 minutes of feeding by ~20 mg/dL (p < 0.01)
despite comparable cumulative food intakes in both groups (Figure 12C). Hence, these results
from genetic loss-of-function experiments further confirm that the activation of duodenal CCK-
A receptors physiologically regulate glucose homeostasis.
Duodenal CCK fails to lower glucose production in response to
high-fat feeding
Previously our laboratory has demonstrated that duodenal lipids fail to activate the gut-
brain-liver axis to lower glucose production in a model of early-onset diet-induced insulin
resistance. On the other hand, we have demonstrated in the current study that the activation of
CCK-A receptor is downstream of gut lipids to lower glucose production via the gut-brain-liver
axis in normal rodents. Therefore, it is now of interest to determine whether the activation of
CCK-A receptors directly through intraduodenal administration of CCK-8 can restore the
activity of the gut-brain-liver axis to regulate glucose production in the same disease model
(Figure 13A,B). Consistent with the findings above, intraduodenal CCK-8 administration
increased the glucose infusion rate (p < 0.05) (Figure 14A) and lowered glucose production (p <
0.001) (Figure 14B,C) in comparison to saline-treated rats when fed on a regular chow diet. In
contrast, intraduodenal CCK-8 administration had no effects on glucose infusion rate (Figure
14A) and glucose production (Figure 14B,C) in comparison to saline-treated rats when fed on a
57
high-fat diet for 3 days. In all treatment groups, glucose uptake remained unchanged (Figure
14D). Hence, these data suggest that high-fat feeding induces duodenal CCK-resistance, leading
to a disruption in the regulation of glucose homeostasis.
58
Figure 3 Schematic representation of the working hypothesis – duodenal CCK activates CCK-A receptors to regulate glucose production and experimental design (pharmacological approach). (A) Proposed model for a duodenal CCK-A receptor pathway to lower hepatic glucose production. CCK-8 is an activator of CCK-A receptors and MK-329 is an antagonist of CCK-A receptors that prevents the activation of the receptors. Intraduodenal administration of CCK-8 lowers glucose production while MK-329 abolishes the effect. (B) Schematic representation of experimental design: intravenous, intra-arterial and intraduodenal catheters were implanted on male Sprague Dawley rats (~250 – 300g). Rats were given 7 recovery days until clamp studies upon which intraduodenal infusion of saline, CCK-8 ± MK-329 were given.
clamp
Intraduodenal SAL or
CCK-8 ± MK-329
Day 1 7
0
90 150 200
[3-3H]-Glucose (0.4 μCi/min)
Insulin (0.8 mU/kg/min)
SRIF (3 μg/kg/min)
Glucose (as needed)
vascular & duodenal catheters
B
A CCK-AR
SAL or CCK-8 ± MK-329
Vagal afferent
MK-329
CCK-8
GP
59
Figure 4 Duodenal CCK activates CCK-A receptors to suppress glucose production (A and B) During the pancreatic clamp (180-200 min), intraduodenal CCK-8 (35 pmol/kg/min; n = 6) infusion increased the glucose infusion rate required to maintain euglycemia (A; *p < 0.05 versus SAL [n = 7]) which was associated with a reduction in glucose production (B; *p < 0.001 versus SAL, MK-329 [n = 7] and CCK-8 + MK-329 [n = 6]). In contrast, coinfusion with MK-329 abolished the effects of CCK-8 on glucose infusion rate (A) and glucose production (B). (C) Suppression of glucose production during the clamp period (180-200 min) expressed as the percentage reduction from basal steady state (60-90 min) glucose production. (*p < 0.01 versus all other groups ). (D) Glucose uptake was unchanged in all groups. Values are shown as mean ± SEM.
A *
*
*
B
C D
60
Figure 5 Schematic representation of the working hypothesis – duodenal CCK activates CCK-A receptors to regulate glucose production and experimental design (molecular approach). (A) Proposed model for a duodenal CCK-A receptor pathway to lower hepatic glucose production. CCK-8 is an activator of CCK-A receptors. OLETF rats are CCK-A receptor deficient such that intraduodenal administration of CCK-8 fails to lower glucose production. (B) Schematic representation of experimental design: intravenous, intra-arterial and intraduodenal catheters were implanted on male LETO and OLETF rats (~250 – 300g). Rats were given 7 recovery days until clamp studies upon which intraduodenal infusion of saline or CCK-8 were given.
CCK-8
CCK-AR
GP
SAL or CCK-8
Vagal afferent
CCK-AR deficient rat (OLETF)
clamp
Intraduodenal SAL or CCK-8
Day 1 7
0
90 150 200
[3-3H]-Glucose (0.4 μCi/min)
Insulin (0.8 mU/kg/min)
SRIF (3 μg/kg/min)
Glucose (as needed)
vascular & duodenal catheters
B
A
61
Figure 6 Duodenal CCK suppresses glucose production in LETO but not in CCK-A receptor deficient OLETF rats (A and B) The body weight of OLETF rats (n = 19) was kept at the same as LETO rats (n = 20) through pair-feeding. (C and D) Intraduodenal CCK-8 infusion (100 pmol/kg/min) increased the glucose infusion rate required to maintain euglycemia during the clamp in LETO rats but not in OLETF rats (C; *p < 0.01 versus all groups) (LETO: SAL [n = 5], CCK-8 [n = 6]; OLETF: SAL [n = 4], CCK-8 [n = 5]). No changes were observed in the glucose infusion rate (C) and glucose production (D) for OLETF rats in response to intraduodenal CCK-8 infusion. (E) Suppression of glucose production during the clamp period (180-200 min) expressed as the percentage reduction from basal (60-90 min) glucose production (*p < 0.01 versus other groups). (F) Glucose uptake was unchanged in all groups. Values are shown as means ± SEM
A
C
B
D
* *
E F
*
62
Figure 7 Schematic representation of the working hypothesis – duodenal CCK activates a gut-brain-liver axis to suppress glucose production and experimental design (A) Proposed model for a gut-brain-liver axis activated by duodenal CCK to lower hepatic glucose production. CCK-8 is an activator of CCK-A receptors. Tetracaine is a local anesthetic that prevents neuronal activations. MK-801 is a potent NMDA receptor antagonist that prevents the activation of the receptors. A separate group of rats received hepatic vagotomy. Intraduodenal administration of CCK-8 fails to lower glucose production in the presence of tetracaine in the duodenum, MK-801 in the NTS, or rats subjected to a hepatic vagotomy. (B) Schematic representation of experimental design: Stereotaxic surgeries were performed on male SD rats (~250 – 300g) 7 days prior to vascular and duodenal cannulations. Rats were given 7 recovery days after the vascular and duodenal cannulation surgeries until clamp studies upon which intraduodenal infusion of saline or CCK-8 ± tetracaine/NTS MK-801 were given.
CCK-8
CCK-AR
SAL or CCK-8 ± tetracaine
GP
NTS MK-801
B
A
NTS cannula clamp
Day 1 13
Intraduodenal SAL or
CCK-8 ± tetracaine
0
90 150 200
[3-3H]-Glucose (0.4 μCi/min)
Insulin (0.8 mU/kg/min)
SRIF (3 μg/kg/min)
Glucose (as needed)
NTS MK-801
Vascular & duodenal catheters (hepatic vagotomy)
7
63
Figure 8 Duodenal CCK can activate a gut-brain-liver axis to regulate glucose production (A and B) Intraduodenal CCK-8 infusion increased the glucose infusion rate required to maintain euglycemia (A; *p < 0.01 versus other groups) and decreased glucose production (B; *p < 0.01 versus other groups). Rats that received tetracaine, NTS MK-801 or hepatic vagotomy failed to suppress glucose production in response to intradudoenal CCK-8 infusions. (SAL [n = 5], CCK-8 [n = 6], CCK-8 + tetracaine [n = 5], CCK-8 + NTS MK-801 [n = 5], CCK-8 + HVAG [n = 6]). (C) Suppression of glucose production during the clamp period (180-200 min) expressed as the percentage reduction from basal (60-90 min) glucose production (*p < 0.05 versus other groups). (D) Glucose uptake was unchanged in all groups. Intraduodenal tetracaine (n = 7), intra-NTS MK-801 (n = 5) or HVAG (n = 6) alone did not affect glucose kinetics. Values are shown as means ± SEM
A B
C D
*
*
*
64
Figure 9 Schematic representation of the working hypothesis – duodenal CCK signaling is downstream of lipid-sensing to regulate glucose production and experimental design (A) Proposed model for the necessity of CCK-A receptor activation in response to duodenal lipids to lower glucose production. MK-329 is a CCK-A receptor blocker that prevents the activation of the receptors. Intraduodenal administration of Intralipids fails to lower glucose production in the presence of MK-329 in the duodenum. (B) Schematic representation of experimental design: vascular and duodenal catheters were implanted on male SD rats (~250 – 300g). Rats were given 7 recovery days until clamp studies upon which intraduodenal infusion of vehicle or Intralipids ± MK-329 were given.
CCK
CCK-AR
GP
lipid
clamp
Intraduodenal SAL or lipid ± MK-329
Day 1 7
0
90 150 200
[3-3H]-Glucose (0.4 μCi/min)
Insulin (0.8 mU/kg/min)
SRIF (3 μg/kg/min)
Glucose (as needed)
vascular & duodenal catheters
B
A
VEH or Lipid ± MK-329
65
Figure 10 Duodenal CCK-A receptor activation is required for lipids to lower glucose production (A and B) Intraduodenal lipids increased the glucose infusion rate (A, *p < 0.01 versus all other groups), and decreased glucose production rate (B, *p < 0.001 versus all other groups) required to maintain euglycemia. The presence of MK-329 abolished the ability of duodenal lipids to suppress glucose production. (VEH; saline + MK-329 alone [n = 11], lipid [n = 10], lipid + MK-329 [n = 7]). (C) Suppression of glucose production during the clamp period (180-200 min) expressed as the percentage reduction from basal (60-90 min) glucose production (*p < 0.01 versus other groups). (D) Glucose uptake was unchanged in all groups. Values are shown as means ± SEM
A B
C D
*
*
*
66
Figure 11 Pharmacological inhibition of CCK-A receptors in the gut disrupts glucose homeostasis during refeeding (A) Schematic representation of the experimental design. Duodenal cannulation was performed on male SD rats (~250 – 280g). Rats were given 5 recovery days until they were subjected to a 40-hr fast. Ten minutes prior to the completion of the fast, an intraduodenal administration of either saline or MK-329 was initiated. Rats were refed on regular chow ad libitum at time 0 min, and food intake and blood glucose levels were monitored at -10, 0, 10 and 20 min. (B) Prior to refeeding, plasma glucose levels were comparable in all rats. After 20 min of refeeding, rats that received MK-329 had significantly higher plasma glucose levels compared to those that received saline. (C) Cumulative food intake was comparable in all groups. Saline (n = 7), MK-329 (n = 6). * p < 0.01 versus saline. Values are shown as means ± SEM
Refed
Day 1 Day 7 at 9am
-10 min 0 60
Intraduodenal SAL or MK-329
Day 5 at 5pm
Fasted
Refed
SD rats
Duodenal catheter
A
B C *
67
Figure 12 Molecular inhibition of CCK-A receptors in the gut disrupts glucose homeostasis during refeeding (A) Schematic representation of the experimental design. Rats were were subjected to a 40-hr fast followed by refeeding. Rats were refed on regular chow ad libitum at time 0 min where food intake and blood glucose levels were monitored at 0, 10 and 20 min. (B) Prior to refeeding, plasma glucose levels were comparable in all rats. After 20 min of refeeding, CCK-A receptor deficient OLETF rats had significantly higher plasma glucose levels compared to LETO rats. (C) Cumulative food intake was comparable in all groups. LETO (n = 6), OLETF (n = 9). * p < 0.01 versus LETO. Values are shown as means ± SEM
Refed
Day 1 at 5pm Day 3 at 9am
0 min 60
Refed
LETO/OLETF rats
fasted
B C
A
*
68
Figure 13 Schematic representation of the working hypothesis – duodenal CCK fails to suppress glucose production in response to high fat feeding and experimental design (A) Proposed model for determining whether duodenal can regulate glucose production in response to 3 days of high fat feeding. After being placed on a lard-enriched high fat diet, intraduodenal administration of CCK-8 fails to lower glucose production. (B) Schematic representation of experimental design: vascular and duodenal catheters were implanted on male SD rats (~250 – 300g). Rats were placed on the regular chow for 4 days, which was replaced by high-fat diet for three days until clamp studies upon which intraduodenal infusion of saline ± CCK-8 were given.
CCK-8
CCK-AR
SAL or CCK-8
GP
HFD ?
clamp
Intraduodenal SAL ± CCK-8
Day 1 7
0
90 150 200
[3-3H]-Glucose (0.4 μCi/min)
Insulin (0.8 mU/kg/min)
SRIF (3 μg/kg/min)
Glucose (as needed)
vascular & duodenal catheters
HFD
4 B
A
69
Figure 14 Duodenal CCK fails to suppress glucose production following 3-days of high-fat feeding (A and B) Intraduodenal CCK-8 administration increased the glucose infusion rate (A, * p < 0.01 versus control), and decreased glucose production rate (B, * p < 0.001 versus control) in rats fed with regular chow (RC). Rats placed on a high-fat diet (HFD) for 3 days failed to respond to intraduodenal CCK-8 infusion to increase the glucose infusion rate (A) and glucose production (B) compared to control. RC: SAL (n = 6), CCK-8 (n = 6); HFD: SAL (n = 4), CCK-8 (n = 5). (C) Suppression of glucose production during the clamp period (180-200 min) expressed as the percentage reduction from basal (60-90 min) glucose production (*p < 0.01 versus other groups). (D) Glucose uptake was unchanged in all groups. Values are shown as means ± SEM
*
*
*
RC HFD RC HFD
RC HFD
RC HFD
A B
C D
70
Basal Insulin (ng/mL) 0.85 ± 0.14 Glucose (mM)
7.6 ± 0.2
SAL CCK-8 Clamp Insulin (ng/mL) 0.73 ± 0.09 0.63 ± 0.13 Glucose (mM) 7.3 ± 0.2 7.2 ± 0.5
Table 2 Plasma insulin and glucose concentrations for SAL and CCK8 during basal and clamp conditions Data are expressed as means ± SEM.
71
5 Discussion
In this study, we revealed a neuronal mechanism by which duodenal CCK can regulate glucose
homeostasis. Duodenal CCK can activate a gut-brain-liver axis through the mediation by CCK-A
receptors and suppress glucose production without affecting glucose uptake. Importantly, this
CCK/CCK-A receptor signaling pathway is essential for duodenal lipids to suppress glucose
production. While these studies were performed in clamped settings, it allowed us to assess the
role of CCK on glucose homeostasis independent of changes in insulin secretions. Moreover, our
intraduodenal administration approach also eliminated the possible interfering effects of gastric
emptying. In a set of fasting-refeeding experiments, we demonstrated that this neuronal network
may also play a role in regulating prandial glucose levels in physiological settings. Consistently,
it has been observed that CCK-A receptor-deficient rats develop an elevated rate of hepatic
glucose production, diabetes and obesity [231-233]. Together, these findings strengthen the
physiological relevance of our findings that gut CCK controls glucose production through a
neuronal network. Here we uncovered a novel neural mechanism for CCK to regulate glucose
homeostasis. Our results suggest that the CCK may be involved in the first line of defense by the
body to adapt appropriately to the incoming nutrients through interactions between the peripheral
nervous system and the central nervous system. Also, in combination with the recent discoveries
that circulating peptide hormones such as insulin or leptin trigger signaling cascades in the brain
to lower glucose production and regulate glucose homeostasis [245-247], we propose that the
central nervous system plays a vital role to convey peptide hormonal signals initiated in the gut
or brain to regulate hepatic glucose production and maintain glucose homeostasis.
72
We showed that duodenal CCK can activate a gut-brain-liver axis to lower glucose production as
an adaptive response to incoming nutrients, the identification of the gut-brain-liver axis was
based on the presence of a gut-brain axis for energy homeostasis by upper intestinal lipids, and a
brain-liver axis for glucose homeostasis by hypothalamic fatty acids [197]. As mentioned in the
introduction, as lipids enter the duodenum, lipases release LCFAs through fat hydrolysis. The
accumulation of LCFAs leads to the stimulation of the brain through the vagus nerve resulting in
inhibition of food intake, thereby regulating energy homeostasis. On the other hand,
intrahypothalamic administration of LCFA led to a suppression of glucose production [248]. As
LCFA accumulates in the hypothalamus, it is esterified by acyl-coenzyme A synthetase to
become LCFA-CoA. The accumulation of LCFA-CoA then leads to the activation of KATP-
channels [248]. Eventually, the hepatic branch of the vagus nerve is stimulated to suppress
glucose production, suggesting that there is direct communication between the brain and the liver
through the vagus nerve. Based on these two neuronal axes, it was first tested whether upper
intestinal lipids can activate a neuronal network to suppress glucose production. Indeed,
duodenal lipids were able to suppress glucose production through a gut-brain-liver axis [20].
Given that CCK is secreted in response to duodenal lipids and we demonstrated that CCK-A
receptor signaling is required for duodenal lipid-sensing, we postulate that CCK is released in
response to duodenal lipids to trigger a CCK/CCK-A receptor signaling mechanism. Since CCK-
A receptors have been located on vagal afferents [249;250] and vagal afferents have been found
to be essential in duodenal lipid-sensing [251], it is probable that the activation of duodenal
CCK-A receptors can lead to stimulation of vagal afferents to send signals to the central nervous
system. One of the brain regions that receive signals from the gut is the hindbrain region NTS
and then the NTS sends signals back to the periphery through the hepatic vagus nerve to suppress
73
glucose production. Hence, our findings also further the fact that the central nervous system can
receive nutrient signals from the gut to regulate glucose homeostasis.
Here, we highlighted the role of the NTS as an integration center in the central nervous system
receiving signals from the peripheral nervous system and sending signals back to the periphery
through the vagus nerve to suppress hepatic glucose production. It has been known that
peripheral CCK can activate CCK-A receptors, leading to the stimulation of the NTS through the
vagus nerve [252-254]. In this study, we specifically showed that duodenal CCK regulates
glucose production through stimulating the NTS via NMDA receptors. As mentioned previously,
the vagus nerve is proposed to be involved in the link between duodenal lipid-sensing and NTS
activation. While peripheral CCK is known to inhibit food intake through activating the NTS, it
has recently been demonstrated that this activation is mediated by the NR2 subunit of NMDA
receptors [255]. NMDA receptors exist as heteromeric complexes composed of NR1, NR2 and
NR3 subunits while MK-801 is only selective for NR1 and NR2 subunits. Since the NR1
subunits are necessary for the NMDA-receptor ion channels to operate, we postulate that the
NR1 subunit should be essential in mediating the effect seen in our study. On the other hand, the
principal neurotransmitter released from vagal afferent terminals in the NTS is glutamate [214]
and glutamate binds with high affinity to the NR2 subunits of the NMDA receptor [256]. These
findings suggest that the NR1 and NR2 subunits may be mediating the effects seen in our study.
Although we were unable to confirm the subunits of the NMDA receptor mediating the effects
seen in our study, we have nonetheless furthered the role of the NTS in mediating glucose-
regulatory effects elicited by duodenal CCK.
74
While we showed that CCK is downstream of lipid-sensing in the duodenum for the regulation
of glucose homeostasis through the gut-brain-liver axis, it was shown that duodenal lipids fail to
lower glucose production in high-fat fed rodents. Hence, one of the goals of the current study
was to determine whether CCK administration can restore the regulation of glucose homeostasis
by the small intestine in high-fat fed rodents. However, we report that there is an acquired defect
in the CCK-A receptor pathway such that duodenal CCK fails to suppress glucose production in
response to three days of high-fat diet. From these results, we postulate that in addition to hepatic
insulin resistance, duodenal CCK-resistance can develop in response to high-fat feeding. Indeed,
while CCK is known to inhibit food intake in normal conditions, it fails to do so in adaptation to
high-fat diets [257-259] with plasma CCK levels upregulated [260;261]. These evidences
support the notion that there is an acquired defect in duodenal CCK-signaling mechanism in
high-fat fed rodents, and this raises the possibility that the restoration of the CCK-signaling
pathway in the gut can lead to restoration of proper glucose and energy homeostasis in diabetes
and obesity.
75
Limitations:
While our findings provided interesting advancements in the understanding of the control of
glucose homeostasis by the small intestine, our findings are not without limitations.
1.) Although we were able to measure CCK levels in the portal and circulation, we did not
measure the CCK levels in the duodenal tissues. In our preliminary experiments, the
measurement of CCK-8 in duodenal tissues was met with the problem that the level of CCK
was beyond the linear range of the standard curve, deeming the values unreliable. Indeed, it
was found in a previous study that the amount of CCK in rat intestine mucosa is up to 174
pmol/g [262]. Hence, we will have to identify a proper dilution factor to allow the level of
CCK in our rat duodenal tissues to be within the linear range of the standard curve, until then
we will not be able to conclude whether the CCK levels achieved through our infusion of
CCK-8 in the duodenum is indeed physiological or pharmacological.
2.) For our fasting-refeeding experiments, we demonstrated that the SD rats infused with CCK-
A receptor blocker or the OLETF rats that are deficient in CCK-A receptors had higher
glucose levels in response to refeeding. Based on this observation, we concluded that the
failure to activate the neuronal network is the reason for the higher glucose levels in these
rats and that the regulation of glucose homeostasis through the neuronal network by duodenal
CCK is physiologically relevant. However, as mentioned in the introduction, upon meal
ingestion, a major source of glucose comes from the meal and the rate at which the glucose
appears in the circulation depends on the gastric emptying rate. In turn, CCK-A receptors
have been implicated in the inhibition of gastric emptying and CCK can actually reduce
postprandial hyperglycemia in humans through slowing gastric emptying [58]. Given these
76
observations, it is possible that the higher glucose observed in the OLETF rats or in response
to CCK-A receptor blocker in SD rats during refeeding was due to a lack of inhibition on
gastric emptying compared to saline, thus allowing glucose to be delivered to the duodenum
for faster absorption. Hence, we are unable to conclude that the higher glucose levels in the
rats abolished of its CCK-A receptor signaling is secondary to accelerated gastric emptying.
A separate set of fasting-refeeding experiments with the same conditions would need to be
performed with an emphasis on the measurement of gastric emptying to confirm that the
differences in glucose levels occurred independent of differences in gastric emptying rate.
3.) Another shortcoming of our study is that we cannot rule out the fact that other gut peptides
may be involved in mediating the observed effects of CCK on glucose production. As
mentioned in the introduction, CCK has been implicated in the secretion of other gut
peptides. For instance, CCK is suggested to be involved in the suppression of ghrelin
secretion and stimulation of GLP-1 and PYY. Hence, to determine whether other gut
peptides are involved, the most convenient way would be to administer CCK to the
duodenum of an overexpression model of ghrelin to assess whether the suppression of
ghrelin is required for CCK to exert its effects on glucose homeostasis. On the other hand, we
will have to infuse CCK to the duodenum of a knockout model of GLP-1 or PYY to
determine whether these gut peptides are involved in the effects of duodenal CCK on glucose
production. However, a knockout model of GLP-1 receptor or PYY is only available in mice
[263;264], such that the CCK-induced gut-brain-liver axis will need to be recapitulated in
mice before it can be confirmed whether GLP-1 receptor is involved in mediating the effects.
77
6 Future Directions
Our study has successfully demonstrated that CCK lies downstream of lipid-sensing in the
duodenum, mediating the activation of a gut-brain-liver axis to regulate glucose homeostasis. As
mentioned in the introduction, intraduodenal administration of glucose or proteins can also lead
to the release of CCK [29;56-58]. On the basis of these findings, it will be of interest to
determine whether duodenal glucose or protein is able to activate the gut-brain-liver axis as
lipids to regulate glucose homeostasis, and if so, whether CCK is involved. If these proposals are
true, CCK would present as a convergence point in small intestinal nutrient sensing to regulate
glucose homeostasis. This will be important because we showed that upon high-fat feeding, the
ability of CCK to regulate glucose homeostasis is lost, it would be of interest to determine
whether high carbohydrate or high protein diets can also lead to duodenal CCK-resistance. These
studies will provide insights into how a balanced diet is integral to effective regulation of glucose
homeostasis by small intestinal nutrients.
Moreover, as we showed that high-fat feeding leads to CCK-resistance, an important future
direction would be to characterize the nature of the resistance to uncover possible ways to
overcome it. Resistance can occur at multiple levels of the pathway, including the
downregulation of available hormones or receptors, or a defect in the signaling cascade. Since
CCK is secreted in response to fat intake, a maintenance high-fat diet can potentially lead to the
adaptation of the body to increase the activity of degradation enzymes to prevent excess CCK
signaling. As such, CCK-resistance may be a result of reduced availability of the peptide, leading
to a reduction in its signaling. However, it has indeed been found that rats on a high-fat diet
78
exhibit elevated postprandial CCK concentrations [265], this suggests that increased CCK
degradation may not be the reason for the development of CCK-resistance. Alternatively, the
development of CCK resistance can occur at the level of the receptor. There could be a reduction
in the expression of the CCK-A receptors in the vagal afferents such that there are less CCK-A
receptors available for activation rendering it more difficult to reach the threshold level of
activation required to stimulate the vagal afferent. To confirm whether this is the case, the
protein expression of CCK-A receptors in the duodenum of regular-chow fed rats would need to
be compared with those of high-fat fed rats. Moreover, since CCK-A receptors are G-protein
coupled receptors, they are subjected to desensitization mechanisms including receptor
internalization and phosphorylation [266]. Therefore, another possibility for the development of
CCK resistance is that there is increased sequestration of receptors such that the downstream
signaling events cannot be triggered. To this end, the amount of CCK-A receptors internalized or
phosphorylated will need to be compared between regular-chow fed rats and high-fat fed rats.
Lastly, the development of CCK-resistance may be due to a defect in its downstream signaling
mechanisms. According to exocrine pancreatic secretion studies, which is another extensively
studied area in regards to CCK-A receptor signaling, it has been discovered that PKA [267] and
PLC [268] play significant roles in mediating the actions of CCK-A receptors in the stimulation
of pancreatic secretions. Similarly, CCK can also activate PKA and PLC in central vagal afferent
fibers [269]. On the basis of these findings, it would be of interest to determine whether PKA
and/or PLC are activated in response to CCK-A receptor signaling in the duodenum to suppress
glucose production. Upon verifying their roles in the duodenum, it would be imperative to
determine whether the activation of these pathways can restore the glucose-regulatory function
of the small intestines in high-fat fed rodents.
79
Furthermore, another future direction stemming from this study is to evaluate the possible role of
CCK-58 in the regulation of glucose homeostasis. In this study, we mainly used the CCK-8 form
of CCK for our experiments, yet there are also other forms of CCK in the circulation: CCK-22,
CCK-33, CCK-39 and CCK-58 [270;271]. While CCK-8 is the shortest biologically active form
of CCK for the CCK-A receptors and it is abundant in the circulation, CCK-58 has been
demonstrated to be another major form in dogs, man and rats [272-275]. Interestingly, CCK-58
can also stimulate intestinal afferent nerve fibers via CCK-A receptors [276]. Functionally,
CCK-58 can inhibit food intake in rats [277] similar to CCK-8, but CCK-58 appears to be more
potent and less readily digested by endopeptidase 24:11 than CCK-8 (longer half-life than CCK-
8) [278]. Therefore, it will be of interest to determine whether CCK-58 can also exhibit the same
glucose-regulatory effects as CCK-8 and whether its actions are affected in high-fat fed rodents.
The results from these experiments would contribute to understanding whether the CCK-
resistance demonstrated is specific for shorter forms of CCK or it is of general resistance to
CCK.
80
7 Conclusion
In this thesis, we examined the possible role of CCK in the regulation of glucose homeostasis
through a neuronal network. Specifically, we demonstrated that, in normal rodents (i) duodenal
CCK can suppress glucose production (ii) this effect requires the activation of CCK-A receptors
(iii) the activation of CCK-A receptors triggers a gut-brain-liver axis (iv) CCK/CCK-A receptor
signaling is required for duodenal lipids to suppress glucose production. However, in rats fed on
three days of high fat diet, duodenal CCK could not suppress glucose production, suggesting that
there is an acquired defect in the CCK-A receptor signaling cascade in response to high fat
feeding. Future studies will need to focus on elucidating the downstream signaling mechanisms
of CCK-A receptors to identify possible ways to restore the regulation of glucose homeostasis by
duodenal lipids.
81
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