Upper Small Intestinal Protein Sensing in the Regulation of Glucose Homeostasis

by

Sophie Claire Hamr

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

Department of Physiology University of Toronto

© Copyright by Sophie Hamr (2016)

ii

Upper small intestinal protein sensing in the regulation of glucose

homeostasis

Sophie Claire Hamr

Master of Science

Department of Physiology University of Toronto

2016

Abstract

High-protein diets improve glucose control in both healthy and type 2 diabetic

individuals. Given that upper intestinal lipids trigger a pre-absorptive, gut-brain-liver

neuronal axis to suppress glucose production (GP), it is urgent to assess the potential

glucoregulatory capacity of intestinal protein sensing. We here demonstrate that in

healthy rodents in vivo, upper small intestinal infusion of protein hydrolysate improved

intravenous glucose tolerance. This was due at least in part to insulin-independent

effects on the liver, given that the same infusion suppressed hepatic GP during a basal-

insulin euglycemic clamp. Co-infusion of tetracaine reversed the ability of protein to

improve glucose tolerance, indicating that a neuronal network is initiated at the level of

the gut to regulate glucose homeostasis. Collectively, these findings show for the first

time that pre-absorptive protein sensing in the upper small intestine improves glucose

tolerance by triggering a neuronal network to lower GP.

iii

Acknowledgments

I would like to acknowledge my supervisor, Dr. Tony Lam, as well as my MSc

committee members Drs. Nicola Jones and Thomas Wolever, for their guidance and

support throughout my graduate studies. I would also like to thank all of the past and

present members of the Lam lab for their direction and assistance with my studies and

for their unconditional moral support, particularly Dr. Frank Duca, Dr. Brittany

Rasmussen and Paige Bauer for their work on this project.

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Table of Contents Chapter 1 Introduction ................................................................................................... 1

1.1 Obesity and diabetes ............................................................................................ 1 1.2 Dietary protein and metabolic homeostasis .......................................................... 3 1.3 Intestinal nutrient sensing in the regulation of metabolic homeostasis ................. 7

1.3.1 Control of food intake ............................................................................... 11 1.3.2 Regulation of glucose homeostasis ......................................................... 14 1.3.3 Dysregulation in obesity and diabetes ..................................................... 18

1.4 Intestinal protein sensing .................................................................................... 22 Chapter 2 Hypothesis and Aims ................................................................................. 26 Chapter 3 Materials and Methods ............................................................................... 29

3.1 Animals .............................................................................................................. 29 3.2 Surgical Procedure ........................................................................................... 29

3.2.1 Preparation of cannulae ........................................................................... 29 3.2.2 Cannulation surgery ................................................................................. 30

3.3 Intestinal infusions and treatments ................................................................ 32 3.4 Intravenous glucose tolerance test ................................................................. 33 3.5 Pancreatic (basal insulin) euglycemic clamp ................................................. 34 3.6 Biochemical analysis ........................................................................................ 35

3.6.1 Plasma glucose ........................................................................................ 35 3.6.2 Plasma amino acids ................................................................................. 35 3.6.3 Plasma [3-3H]-glucose specific activity ..................................................... 37

3.7 Tracer-dilution calculations ............................................................................. 37 3.8 Statistical analysis ............................................................................................ 38

Chapter 4 Results ......................................................................................................... 40 Chapter 5 Discussion ................................................................................................... 54 Chapter 6 References ................................................................................................... 65

v

List of Tables

Table 1: Typical amino acid content of Casein enzymatic hydrolysate from Sigma-

Aldrich (Product

22090).................................................................................................51

Table 2: Body weight and cumulative post-surgical food intake of the groups of rats that

underwent either the intravenous glucose tolerance test or clamp

protocol...................52

Table 3: Plasma glucose concentrations during the basal insulin euglycemic

clamp.....53

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List of Figures

Figure 1: Intestinal nutrient sensing in the regulation of metabolic

homeostasis............25

Figure 2: Schematic of the working

hypothesis..............................................................28

Figure 3: Gut infusion-IVGTT protocol to evaluate the effects of upper small intestinal

protein sensing on glucose

tolerance.............................................................................47

Figure 4: Upper small intestinal infusion of protein improves glucose

tolerance............48

Figure 5. Upper small intestinal casein hydrolysate increases the glucose infusion rate

during the basal insulin euglycemic

clamp.....................................................................49

Figure 3: Gut infusion-IVGTT protocol to evaluate the effects of upper small intestinal

protein sensing on glucose

tolerance.............................................................................50

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List of Abbreviations ACS Acyl-CoA synthetase AUC Area under the curve CCK Cholecystokinin CD36 Cluster of differentiation 36 DPP-IV Dipeptidyl Peptidase-4 DVC Dorsal vagal complex EEC Enteroendocrine cell ENS Enteric nervous system FXR Farnesoid X receptor GI Gastrointestinal GLP-1 Glucagon-like peptide 1 GIP Glucose-dependent insulinotropic peptide GIR Glucose infusion rate GPCR G-protein coupled receptor HbA1c Glycated hemoglobin IVGTT Intravenous glucose tolerance test i.p. Intraperitoneal LCFA Long-chain fatty acid NMDA N-methyl-D-aspartate NTS Nucleus of the solitary tract OGTT Oral glucose tolerance test OLETF Otsuka-Long-Evans-Tokushima Fatty PepT1 Peptide transporter 1 PLC Phospholipase C POMC Pro-opiomelanocortin PKA Protein kinase A PKC Protein kinase C PYY Peptide YY Ra Rate of glucose appearance Rd Rate of glucose disappearance SD Sprague Dawley SGLT-1 Sodium-glucose cotransporter 1

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Published Manuscripts that contributed to this thesis

Hamr SC, Wang B, Swartz TD and Duca FA. Does nutrient sensing determine how we

“see” food? Curr Diab Rep 15(6):604 (2015).

Duca FA, Bauer PV, Hamr SC and Lam TK. Glucoregulatory relevance of small

intestinal nutrient sensing in physiology, bariatric surgery and pharmacology. Cell Metab

22(3):367-80 (2015).

Bauer PV, Hamr SC and Duca FA. Regulation of energy balance by a gut-brain axis

and involvement of the gut microbiota. Cell Mol Life Sci 73(4):737-55 (2016).

1

Chapter 1

Introduction

1.1 Obesity and diabetes

Over 700 million individuals are obese, and over 300 million have type 2 diabetes

worldwide, rendering a state of global epidemic1,2. Given that obesity and diabetes are

associated with a number of life-threatening co-morbidities, and collectively cost the

Canadian healthcare system over $20 billion per year3,4, there is an urgent need to

understand their pathologies in hopes to develop successful treatments.

The etiologies of these two diseases are complex and interconnected given that

approximately 80% of type 2 diabetic individuals are overweight or obese5. Obesity and

diabetes are linked to a number of genetic and environmental factors, which collectively

lead to a dysregulation in the homeostatic control of energy balance and glucose

homeostasis. In healthy individuals, energy homeostasis is tightly regulated by a

multitude of interconnected signaling networks, which integrate signals of energy status

and regulate exogenous nutrient intake and endogenous nutrient metabolism,

accordingly. The classical view of glucose regulation and of diabetes development is an

islet-centered view, whereby blood glucose levels are primarily regulated by the

pancreatic hormones insulin and glucagon. Insulin resistance develops at the level of

peripheral tissues, leading to a compensatory increase in pancreatic insulin secretion.

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As insulin resistance worsens, β-cell exhaustion occurs and insulin secretion no longer

compensates, resulting in hyperglycemia and a diagnosis of type 2 diabetes6. However,

50% of glucose tolerance is accounted for by insulin-independent mechanisms7, which

remain incompletely understood. Schwartz et al. recently proposed the importance of a

“brain-centered glucoregulatory system,” whereby the brain senses circulating nutrients

to regulate plasma glucose independently of insulin8. Indeed, nutrient sensing

mechanisms are an important form of insulin-independent regulation on energy

homeostasis.

Interestingly, more recent evidence brings to light the importance of nutrient sensing by

the gastrointestinal (GI) tract, indicating that nutrients generate critical negative

feedback on energy homeostasis upon initial contact with the body9. The impact of GI

or “gut” nutrient sensing on glucose homeostasis is highlighted by the fact that

metformin, the most widely prescribed drug for the treatment of type 2 diabetes, has

been found in both rodents and humans to exert effects on glycaemia via a gut-

localized site of action10,11. Further, when the GI tract is surgically re-arranged to treat

obesity, redirecting nutrient flux and implicating distal intestinal nutrient sensing, a drop

in plasma glucose levels occurs almost immediately, prior to any weight loss12,13.

Finally, pharmacological treatments increasing levels of the nutrient-stimulated gut

hormone glucagon-like peptide 1 (GLP-1) have proven effective in regulating glycaemia

in type 2 diabetic individuals14. While the mechanisms underlying these therapies

remain incompletely understood, they collectively implicate an urgent need to further

3

elucidate intestinal nutrient signaling in the regulation of glucose homeostasis and

pathogenesis of obesity and diabetes.

1.2 Dietary protein and metabolic homeostasis

In comparison to the increasing knowledge of how carbohydrates and lipids regulate

glucose homeostasis, the effects of dietary proteins remain less clear. Numerous

studies show that intravenous infusion of amino acids in a hyperinsulinemic,

euglycemic condition induces acute insulin resistance at the level of glucose uptake by

peripheral tissues15-19. In vitro studies in isolated myocytes indicate that amino acids

inhibit insulin-stimulated glucose uptake likely via competitive inhibition of glucose

oxidation (glycolysis)20-22, activation of mTOR23, and inhibition of the insulin signaling

pathway24. Interestingly, however, these findings are not consistent, given that others

found that elevating plasma amino acids had no effect on glucose uptake during a

hyperinsulinemic clamp in healthy men25, and actually increased glucose uptake to

lower glucose levels during unclamped hyperinsulinemia26. Thus, in order to examine

how dietary protein affects glucose turnover in the context of high-protein feeding,

Rossetti et al pair-fed rats either a high-protein (60% protein) or low-protein (18%

protein) diet27. Following 10 days of dietary intervention, rats fed the high-protein diet

had increased fasting glucose levels and decreased insulin sensitivity during a

hyperinsulinemic clamp at the levels of both hepatic glucose production and peripheral

glucose uptake27. This study suggests that acutely increasing dietary protein intake

may be detrimental to glucose homeostasis, due to amino acid-induced insulin

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resistance. However, as described below, longer dietary interventions in fact show that

high protein feeding improves energy balance and glucose control, suggesting that

dietary proteins may exert other, yet to be identified effects that positively impact

metabolic homeostasis.

High-protein diets have been widely prescribed as a dietary intervention in overweight

individuals, and over a 6-month test period, high-protein diets have more successful

weight loss outcomes than conventional, high-carbohydrate weight-loss diets28-30.

These high-protein diet-treated individuals not only experience a greater overall weight

loss, but also display decreased fat mass and reduced fasting plasma free fatty-acids,

triglycerides and cholesterol29,30, indicating an overall improvement in lipid

homeostasis. These results were confirmed in healthy rodents, where a 42-day high-

protein diet reduced body weight and fat mass accumulation compared to a low-protein

diet31. Interestingly, individuals fed a high-protein weight-loss diet often have

corresponding increases in insulin sensitivity and lowering of glycated hemoglobin

(HbA1c)28,29,32,33, suggesting that increasing dietary proteins can improve glucose

homeostasis in addition to their weight-loss properties. Indeed, when Samaha and

colleagues followed up after 1 year of high-protein versus high-carbohydrate weight-

loss dieting, the high-protein group displayed lower levels HbA1c despite no persisting

effects on body weight34. However, improvements in glucose homeostasis following

long-term high-protein feeding may be secondary to the initial weight-loss, given that

modest weight loss per se is associated with improved glycemic control35. Thus, it is

important to look at the effects of short-term high-protein feeding independent of any

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weight loss. On this note, when subjects were fed a weight-maintenance diet for 5

weeks, a high-protein diet improved glucose tolerance and HbA1c levels compared to a

high-carbohydrate diet, independent of any change in body weight36. Along with an

epidemiological analysis linking higher dietary amino acid intake with a lower risk of

type 2 diabetes37, these findings highlight the importance of looking more closely at the

effects of high-protein feeding to regulate blood glucose.

Paralleling these human studies, rodents fed a high milk-protein diet for 6 months

exhibited lower fasting plasma glucose and improved glucose tolerance as assessed

by an oral glucose tolerance test (OGTT) compared to rodents fed a normal-protein

diet38. However, it is important to note that in this study, rats were fed the diets ad

libitum, resulting in a 13% lower total food intake in the high-protein diet group38.

Indeed, both rodents and humans commonly exhibit lower food intake in response to a

high-protein diet31,33,39. Thus, in a follow up study, rats were pair-fed to look at the

effects of high-protein feeding independent of effects on food intake. In this context,

high-protein feeding no longer decreased fat mass, however the high-protein-fed rats

displayed persistent improvements in glucose tolerance and fasting plasma glucose40.

This study provides strong evidence that dietary protein can regulate glucose

homeostasis independent of effects on food intake and body weight.

In an acute setting, rats fasted for 24 hours and fed either a standard mixed meal or an

equivalent caloric load of protein hydrolysate experienced a much lower glucose

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response to the protein meal41. However, in this case, it is impossible to distinguish the

glucose-lowering effects of the protein per se from the coinciding lack of dietary

carbohydrate contributing to the glucose response. Indeed, this variable must also be

considered in the case of the high-protein diet models described earlier, given that

protein was always substituted for carbohydrates in these diets. Thus, a model to look

at the glucose-lowering effects of protein per se is presented in studies where protein is

administered in addition to a standard meal. In one such study, overnight-fasted human

subjects consumed a standard maltodextrin beverage, with or without the addition of

protein hydrolysates42. The addition of protein lowered the glucose response to the

maltodextrin beverage, despite the equal contribution of carbohydrates and the higher

caloric load42. This phenomenon is mirrored by two similar studies where protein was

administered in the form of a whey-protein beverage either directly or 30 minutes prior

to a standard meal, while control groups did not receive a pre-load. In both cases, pre-

loading with a whey protein bolus lowered the glucose response to the subsequent

meal43,44. It is interesting to note that although amino acids stimulate insulin secretion,

the lower glucose response observed in the study by Akhavan et al was not associated

with higher insulin levels43. Thus, it would seem that the additional dietary protein load

exerts insulin-independent effects on glucose regulation in order to improve glucose

tolerance during the subsequent meal.

Our laboratory recently investigated this insulin-independent, protein-induced regulation

of glucose homeostasis. Healthy, male Sprague-Dawley rats were overnight-fasted and

fed a 19 kcal meal with either 20% or 60% calories from protein (milk-derived casein).

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While the total calorie intake, rate of food intake, and insulin response were indeed

identical between groups, the 60%, high-protein diet resulted in a lower plasma glucose

response from 30-60 minutes post meal initiation45. Interestingly, although the insulin

response was the same, the high-protein meal increased plasma glucagon whereas the

low-protein meal did not. Indeed, glucagon signaling was partially responsible for the

glucose-lowering effects of protein, as hindbrain glucagon receptor blockade reversed

the improvement in glucose response seen at 60 minutes45. This is consistent with the

previously documented glucagon-stimulating effects of protein and glucose-lowering

effects of brain glucagon signaling45-47. However, it is important to note that glucagon

receptor blockade did not reverse the glucose-lowering effects of high-protein feeding

during the earlier time-points, indicating that there are additional, potentially insulin- and

glucagon-independent mechanisms at play. Given that this glucose-lowering response

is robust within just 30 minutes of feeding, one could speculate that negative feedback

mechanisms arise at the level of the gut upon interaction with incoming nutrients. This

theory is in fact not unprecedented given that, as described below, sensing of nutrients

such as protein by the GI tract can initiate rapid and potent negative feedback on

whole-body metabolic homeostasis.

1.3 Intestinal nutrient sensing in the regulation of metabolic homeostasis

The GI tract is the primary site of interaction between incoming nutrients and the body,

providing a site for early negative feedback on metabolic homeostasis. The extensive

8

neural and humoral connectivity of the gut wall to other important energy-regulating

areas, such as the brain, allows it to effectively relay information about incoming

nutrients in order to prevent energy excess upon nutrient absorption into the circulation.

The upper small intestine, where the majority of nutrient absorption occurs, is the major

feedback site responsible for energy regulation48.

Nutrient signaling is mediated by the release of gut peptide hormones from

enteroendocrine cells (EECs). EECs, expressed throughout the gut epithelial mucosa,

projecting their apical surfaces into the intestinal lumen, allowing them to act as

chemoreceptors, detecting nutrients predominantly via cell-surface G-protein coupled

receptor (GPCRs) or solute transporters49. In response to the detection of nutrients,

EECs release gut peptides from their basal surfaces into intestinal capillaries. Thus, the

EEC is proposed as the gut’s nutrient-sensing cell, directly sensing luminal nutrients

and generating humoral signals in response. Gut peptide hormones are thought to be

associated with individual EEC types that differ in the gut peptides they produce and

their localization along the GI tract. The proximal small intestine expresses “I cells,”

which produce the peptide cholecystokinin (CCK), and “K-cells” that produce glucose-

dependent insulinotropic peptide (GIP), while the more distal “L cells” produce GLP-1

and Peptide YY (PYY). However, there is accumulating evidence for co-expression of

gut peptides thought to be produced in distinct cell types, suggesting that the EEC may

in fact be a single cell type that produces varying spectra of peptides depending on its

environment50,51, though this is continually debated52.

9

Specific and independent sensory and signal transduction mechanisms mediate gut

peptide release in response to individual types of nutrients. EECs express a number of

nutrient-specific GPCRs such as the fatty-acid receptors GPR4053,54 and GPR12054,55

and the amino acid and peptide-receptive calcium sensing receptor (CaSR)56, as well

as solute transporters such as sodium-glucose transporter 1 (SGLT-1)57 and peptide

transporter 1 (PepT1)58. Via these chemosensory machineries, luminal nutrients

promote the exocytosis of gut peptides through intracellular metabolism, second-

messenger signaling cascades and/or membrane depolarization59. Indeed, chemical

inhibition or genetic knockout of one of these nutrient-sensing receptors or transporters

will results in impaired CCK, GLP-1 or GIP release in response to feeding53,60-63.

However, refuting this classical model of EEC nutrient sensing, there is accumulating

evidence for indirect mechanisms through which nutrients may alternatively stimulate

gut peptide release. For example, numerous studies show that lipid absorption and

packaging into chylomicrons, and the subsequent release of Apolipoprotein A-IV from

enterocytes, is required for dietary lipids to stimulate the release of CCK and GIP64-66.

Bile acids, components of bile that are released form the liver into the intestinal lumen

in response to nutrient ingestion to aid with the solubilization and digestion of lipids,

have also been implicated in stimulating gut peptide release. Bile acids themselves

may act directly on EECs to regulate gut peptide secretion via their endogenous

receptor, the farnesoid X receptor (FXR), though the evidence to date is

contradictory67,68. Nonetheless, whether nutrients signal directly on or within EECs, or

indirectly via other cell types, the resulting release of gut peptide hormones is the first

step in the initiation of nutrient-induced negative feedback axes.

10

Upon exocytosis from EECs, gut peptides are secreted into the portal circulation to act

directly on target organs in an endocrine fashion. For example, systemic plasma CCK

levels increase in the postprandial state in both humans and rodents69,70, and the brain

expresses CCK receptors that are implicated in metabolic regulation71. The brain also

expresses GLP-1 receptors72,73, however unlike CCK, GLP-1 is rapidly degraded in

circulation by the enzyme Dipeptidyl Peptidase-4 (DPP-IV), resulting in less than 10%

of GLP-1 reaching the systemic circulation following secretion74. GLP-1 may instead

reach the brain via lymph vessels, given that in the postprandial state, lower levels of

DPP-IV and higher levels of GLP-1 are found in the lymph than in the portal vein75-77.

The gut peptide PYY is likely to signal predominantly in an endocrine fashion, given

that circulating PYY levels peak 1.5 hours after feeding and remain elevated for several

hours78. Alternatively, CCK and GLP-1, which display more transient peaks, may in fact

be more important in paracrine signaling with the gut.

The proximal small intestine is densely innervated by the vagus nerve, with afferent

fibers (projecting to the brain) far outnumbering efferent fibers79,80. Extensive evidence

indicates that the acute regulation of food intake and plasma glucose in response to

intestinal nutrients is mediated by these vagal afferents. Indeed, vagal afferents

express receptors for CCK, GLP-1 and PYY, and activation of these vagal gut peptide

receptors leads to afferent firing81. Vagal afferent dendrites innervate the lamina propria

of the gut mucosa, rather than EECs directly, indicating that gut peptides likely diffuse

to nearby vagal targets in a paracrine fashion once released81. Alternatively, neurons of

the enteric nervous system (ENS), which also express gut peptide receptors and are

11

situated in between EECs and vagal afferents, may act as an intermediate neuronal

relay82-84. One study found that the nutrient-sensing receptor GPR40 is expressed by

vagal afferent neurons themselves, suggesting that in some cases nutrients diffuse

through the gut epithelium to activate vagal afferents directly85. Nonetheless, nutrient

influx results in the activation and firing of vagal afferent neurons, which subsequently

relay a signal to the nucleus of the solitary tract (NTS) in the dorsal vagal complex

(DVC) of the hindbrain, where vagal afferents terminate86. Indeed, intraduodenal

nutrient infusion leads to c-fos activation of NTS neurons, and this response is blocked

by treatment of gut vagal afferents with the neurotoxin capsaicin87. Gut-derived signals

are integrated by the NTS, which relays to other brain areas to regulate metabolic

homeostasis accordingly88. This EECàvagal afferentàhindbrain axis mediates a wide

range of feedback systems activated by nutrients in the intestine, including important

regulation on food intake and glucose homeostasis (Figure 1).

1.3.1 Control of food intake

Fasted rats decrease their rate of feeding within minutes of meal initiation89, indicating

the presence of negative feedback on food intake before nutrients reach the circulation

(the term “pre-absorptive” hereafter refers to effects generated by nutrients prior to the

release of nutrients into the portal and systemic circulation, but not necessarily prior to

absorption by enterocytes). This pre-absorptive feedback is initiated within the small

intestine, given that a suppression of food intake is entirely absent during sham

feeding, when food is drained from gastric fistulas, never reaching the duodenum90.

12

Alternatively, infusion of a nutrient solution directly into the lumen of the duodenum

results in a rapid suppression of sham feeding91. Surgical ablation of the

subdiaphragmatic vagus or gut capsaicin treatment completely reverses the

suppressive effects of intraduodenal proteins, carbohydrates or lipids on sham

feeding92,93. Likewise, capsaicin treatment of the hindbrain blocks the effect of intestinal

nutrients to suppress sham feeding94. Together, these results indicate that the acute

feedback on food intake in response to duodenal nutrients is entirely mediated by a

vagal gutàhindbrain neuronal axis.

CCK, often referred to as the “satiety hormone,” has long been associated with acute

control of food intake in both rodents and humans95,96. In humans, intravenous infusion

of a CCK receptor antagonist delays feelings of “fullness” during a meal97, and reverses

the suppression of ad libitum food intake by intraduodenal lipid infusion98. In rodents,

CCK receptor knockout, or co-infusion of a CCK receptor antagonist, reverse vagal

afferent firing and suppression of food intake in response to intraduodenal nutrients99-

101. Likewise, peripheral (intraperitoneal; i.p.) CCK-8 (the biologically active form of

CCK) injection activates brainstem neurons to suppress feeding, and this effect is

completely abolished by intestinal capsaicin, subdiaphragmatic vagotomy, or hindbrain

capsaicin94,102,103. This CCK-mediated gutàbrainstem axis requires CCK-A (CCK-1),

but not CCK-B receptors, which are indeed expressed by vagal afferent fibers101,104,105.

Together, these results suggest that CCK produced by the gut regulates satiety entirely

via vagal CCK-1 receptors. While CCK receptors are expressed in the brain, and

intracerebroventricular CCK administration lowers feeding106, central receptors may

13

mediate pathways by centrally-produced, and not intestinally-produced CCK103, given

that central CCK-1 receptor inhibition does not block feeding suppression by intestinal

nutrients107.

Postprandial GLP-1 levels correlate with lower hunger scores in humans108, suggesting

a role for GLP-1, in addition to CCK, in the suppression of appetite. Indeed, in rodents,

i.p. GLP-1 injection suppresses food intake, and this effect is blocked by vagotomy or

capsaicin, implicating a parallel role for vagal afferent GLP-1 action in nutrient-induced

satiety109,110. In humans, intravenous GLP-1 infusion dose-dependently suppresses

energy intake during an ad libitum meal111. Patients that have undergone truncal

vagotomy surgery as a treatment for duodenal ulcers provide a comparative human

model to subdiaphragmatic vagotomized rodents, and these individuals are

characterized by impaired suppression of feeding in response to intravenous GLP-1

infusion112. Thus, there is extensive evidence that like CCK, GLP-1 released in

response to luminal nutrients acts on local, vagal afferents to trigger a gut-brain axis to

suppress feeding.

At the level of the hindbrain, the activation of NTS neurons by vagal afferents and

subsequent neuronal relay is incompletely understood, but involves N-methyl-D-

aspartate (NDMA) receptor transmission and subsequent phosphorylation of ERK 1/2

and synapsin I within the NTS113-115. The distinct neuronal populations within the NTS

that are activated by vagal afferent signaling appear to include anorexigenic pro-

14

opiomelanocortin (POMC)-expressing neurons116. Indeed, blockade of NTS

melanocortin receptor (MC4R) signaling reverses the activation of NTS ERK1/2 and

suppression of feeding in response to peripheral CCK117. NTS neurons then project to

other brainstem nuclei which contain neuronal motor output that controls the

behaviours involved in feeding, supporting a possible brainstem-isolated reflex to

control food intake. Indeed, intestinal nutrients suppress food intake in decerebrate rats

with only the brainstem intact118. However, NTS neurons also project to many higher-

order brain regions involved in the control of feeding, including the hypothalamic

arcuate and paraventricular nuclei. Indeed, Otsuka-Long-Evans-Tokushima Fatty

(OLETF) rats, which lack the CCK-1 receptor and are subsequently hyperphagic and

mildly obese, display altered hypothalamic melanocortin neuropeptides119, supporting a

role for the hypothalamus in regulation of feeding by gut peptides.

1.3.2 Regulation of glucose homeostasis

While the suppression of food intake by gut peptide-mediated nutrient sensing

mechanisms has been studied extensively, far less is known about the role of nutrient

signaling in the regulation of blood glucose. However, accumulating evidence supports

a role for the GI tract in acutely regulating whole-body glucose metabolism. The first

observation that the gut could regulate blood glucose was the discovery of the “incretin”

effect, whereby oral glucose administration results in roughly double the insulin

response compared to delivery of the same amount of glucose intravenously120.

Indeed, the gut peptides GIP and GLP-1, both secreted in response to intestinal

15

nutrients, act directly on pancreatic β-cells to stimulate glucose-dependent insulin

secretion, subsequently improving glucose tolerance121-123. Interestingly, when GLP-1

is infused intravenously, glucose tolerance improves not only through an increase in

insulin secretion but also by insulin-independent increases in glucose disposal124. In

another study, intravenous GLP-1 infusion during a basal insulin pancreatic clamp

suppressed hepatic glucose production independent of any pancreatic action125.

Although the mechanisms are unknown, these results suggest that the glucoregulatory

capacity of gut-derived signals is not in fact limited to the incretin effect. Brain GLP-1

signaling may play a role, given that central GLP-1 infusion stimulates muscle glucose

uptake independent of muscle insulin receptors126, and hypothalamic GLP-1 infusion

suppresses hepatic glucose production127. These studies suggest that gut-derived

GLP-1 may reach the brain to signal via central GLP-1 receptors to regulate blood

glucose, though again, this signaling axis could be more relevant to centrally-produced

GLP-1.

Given the implication of vagal afferent gut peptide receptors in the regulation of food

intake, vagal GLP-1 receptor signaling within the gut might also play a role in the

regulation of glucose homeostasis by GLP-1. To investigate this hypothesis, a rat

model was developed with specific knockdown of GLP-1 receptors in vagal afferent

neurons through delivery of an shRNA lentiviral vector into the nodose ganglia128. In

this rat model, a 50% knockdown of vagal afferent GLP-1 receptors resulted in

increased post-meal glucose levels following an overnight fast128, confirming that

paracrine, vagal GLP-1 signaling plays a physiological role in glucose regulation.

16

Furthermore, a low oral dose of DPP-IV inhibitor, which selectively inhibits intestinal

DPP-IV activity, increases vagal afferent firing and improves glucose tolerance, and

this effect is absent in GLP-1-/- mice129, further supporting the role of vagal GLP-1

signaling within the intestine to regulate glucose homeostasis. Of note, in addition to

innervating the gut wall, vagal afferent fibers innervate the portal vein and this is a

second site at which nutrients may trigger a gut-brain axis prior to reaching the

systemic circulation. Indeed, infusion of glucose directly into the portal vein increases

whole-body GU to lower plasma glucose, and this is blocked by co-infusion of a GLP-1

receptor antagonist130, implicating a potential role for hepatoportal GLP-1 receptors in

regulating whole-body glucose homeostasis via a gut-brain axis. Nevertheless, further

work is required to dissect intestinal and hepatoportal GLP-1 signaling in the regulation

of glucose homeostasis via a gut-brain axis.

Gut peptide regulation of glucose homeostasis is in fact not limited to the well-known

“incretin hormones” GLP-1 and GIP, as intravenous infusion of CCK increases insulin

secretion and lowers plasma glucose levels131. Indeed, β-cells express the CCK

receptor132, and CCK-/- mice display impaired insulin response to i.p. glucose

injection133. In vitro studies show that like GLP-1 and GIP, CCK stimulates insulin

secretion from β-cells by enhancing GSIS134-136. In addition, CCK-1 receptor signaling

within the intestine contributes to the glucoregulatory effects of CCK. Intraduodenal

infusion of a CCK-1 receptor inhibitor dysregulates the glucose response to mixed-meal

feeding in healthy rodents137, indicating that intestinal CCK-1 receptor signaling is in

fact required for normal glucose homeostasis during digestion.

17

In order to dissect the mechanisms through which intestinal nutrient sensing might

regulate glucose homeostasis via CCK, we examined the effect of intraduodenal

nutrient infusion on glucose kinetics during a basal insulin euglycemic pancreatic

clamp. During the clamp, intraduodenal infusion of Intralipid (a lipid emulsion) rapidly

and potently suppressed hepatic glucose production, but did not alter glucose

uptake138. This effect on glucose production was mediated by a gut-brain-liver neuronal

axis, involving vagal afferents, NMDA receptor transmission in the NTS, and the

hepatic vagal efferent138. At the level of the gut, the effect of Intralipid to lower glucose

production required esterification of long-chain fatty acids (LCFA) to LCFA-CoA by

acyl-CoA synthetase (ACS), implicating cellular uptake and subsequent metabolism of

fatty-acids, rather than activation of cell-surface receptors, in the stimulation of gut

peptide release. Co-infusion of a CCK-1 receptor antagonist or genetic knockout of the

CCK-1 receptor reversed the effects of intraduodenal lipids to suppress glucose

production, while intraduodenal infusion of CCK-8 per se suppressed glucose

production137, indicating that a release of CCK in response to intestinal LCFA-CoA

mediates this gut-brain-liver axis. Blocking duodenal PKC-δ negated the ability of

intraduodenal lipids to lower glucose production139, indicating that LCFA-CoA

stimulates the release of CCK via activation of PKC-δ, a signaling pathway that has

been confirmed in vitro140,141. Of note, intraduodenal infusion of a PKC-δ inhibitor

resulted in a dysregulation of glucose response during an un-clamped feeding

paradigm, proving the physiological relevance of this duodenal lipid-sensing pathway.

The CCK-1 receptor, a GPCR, activates both phospholipase C (PLC)- and protein

kinase A (PKA)-dependent pathways, both of which have been shown to be necessary

18

for activation of vagal afferent neurons by CCK142. During the clamp, activation of PKA

was required for intraduodenal lipids or CCK-8 to stimulate vagal afferent firing and

suppress hepatic glucose production143. Thus, a LCFAàLCFA-CoAàPKC-δ

àCCKàCCK1RàPKA signaling axis within the upper small intestine activates a gut-

brain-liver neuronal axis to suppress hepatic glucose production and maintain glucose

homeostasis in healthy rodents in vivo144. This confirms that in parallel to a gut-brain

axis to regulate food intake, pre-absorptive lipids within the intestine activate a neuronal

gut-brain-liver axis to regulate glucose homeostasis (Figure 1). Whether a similar

negative feedback axis to regulate glucose homeostasis is initiated by upper intestinal

carbohydrates or protein remains to be investigated.

1.3.3 Dysregulation in obesity and diabetes

Obesity, subsequent dysregulation of glucose homeostasis and the onset of diabetes

develop from a combination of genetic and environmental factors that disrupt that

homeostatic control of food intake and glucose homeostasis. One such environmental

factor is the overconsumption of a highly palatable “western diet” that is high in sugar

and fat. Evidence suggests that the consumption of a high-fat diet alters intestinal

nutrient-sensing pathways that, as described above, are important factors in the

regulation of energy homeostasis, and that this may in fact be an important contributor

to the development of obesity and diabetes. Understanding the precise mechanisms of

this disruption to nutrient sensing has the potential to unveil novel therapeutic targets to

restore homeostasis in disease.

19

Following 21 days of high-fat diet feeding in rats, the reduction of food intake in

response to intraduodenal fatty acid infusion is attenuated compared to rats maintained

on a low-fat diet145. Likewise, humans maintained on a high-fat diet for just 14 days are

no longer sensitive to the satiating properties of intraduodenal lipid infusion146. In

another study, 14 day high-fat diet-fed humans consuming an ad libitum meal reported

lower feelings of “fullness” and ate more than when fed a balanced diet for 14 days

beforehand147. Thus, it is evident that high-fat feeding leads to a disruption in intestinal

nutrient sensing mechanisms, at least in the regulation of food intake, but it is unclear

which precise mechanism(s) in the feedback axis might be disrupted.

There is evidence that high-fat feeding alters the expression of the GPCR’s and

transporters through which gut epithelial cells sense luminal nutrients. EECs isolated

from the small intestines of high-fat diet-fed mice have lower expression of the nutrient-

sensing transporters SGLT-1 and PepT1 as well as the fatty-acid receptor GPR120

compared to those of healthy mice148. Likewise, overweight and obese humans have

lower duodenal expression of the fatty-acid receptor GPR11954. However, in the same

study, the authors identified higher expression of duodenal cluster of differentiation 36

(CD36; a lipid transporter implicated in gut peptide release) and GPR12054,149. This is

consistent with another study in rodents, where rats fed a high-fat diet for 10 weeks

displayed higher expression of the fatty-acid receptors GPR40, GPR41 and

GPR120150. Increased fatty-acid receptor expression suggests that the disruption in

20

nutrient sensing may occur downstream of lipid receptor activation, leading to a

compensatory increase in receptor expression.

Indeed, the same obese individuals who showed increased duodenal CD36 and

GPR120 displayed decreased numbers of CCK- and GLP-1-expressing cells54.

Similarly, rats fed a high-fat diet for 10-weeks displayed lower intestinal CCK and GLP-

1 protein expression150. This suggests that high-fat feeding can lead to a decrease in

CCK and GLP-1 production, which may at least in part contribute to the disruption in

the ability of nutrient sensing to suppress food intake described above. However, of

note, the impaired satiation by oral or intraduodenal nutrients in 14-day high-fat diet-fed

humans was not in fact associated with lower release of CCK or GLP-1146,147. This

suggests that in high-fat feeding, even if nutrients are able to stimulate the release of

sufficient CCK or GLP-1, the ability of gut hormones to initiate the gut-brain feedback

axis may be impaired.

Indeed, rats fed a high-fat diet for 3 weeks do not suppress food intake in response to

i.p. CCK-8 injection at a dose that significantly reduces food intake in low-fat diet-fed

rats151. Likewise, we found that in rats fed a 3-day high-fat diet, neither intraduodenal

Intralipid nor CCK-8 infusion suppresses hepatic glucose production during the

pancreatic clamp137,138. The inability of CCK to initiative negative feedback on food

intake or glucose production implicates a potential role for resistance at the level of

vagal afferent gut peptide receptors. Indeed, 3-week high-fat diet-fed rats display

21

reduced c-fos activation of neurons in both the nodose ganglion (vagal afferents) and

the NTS in response to i.p. CCK injection or intraduodenal lipid infusion152-154,

implicating reduced neuronal sensitivity to CCK. We found that the resistance for CCK

to suppress hepatic glucose production occurred at the level of the activation of PKA

and subsequently vagal afferent firing by intraduodenal CCK143. As such, bypassing

CCK receptor activation and infusing a PKA activator directly into the duodenum

restored vagal afferent firing and suppression of hepatic glucose production143. Thus,

through unclear mechanisms, high-fat feeding disrupts the ability of CCK activation of

the CCK-1 receptor to stimulate PKA signaling within the gut (Figure 1). More work is

required to understand the precise mechanisms through which high-fat feeding disrupts

the gut-brain axis to regulate various physiological functions, and the potential to

restore intestinal nutrient signaling pathways to treat obesity and diabetes.

Evidently, intestinal nutrient sensing plays an important role in the acute regulation of

both food intake and blood glucose during feeding, and these regulatory mechanisms

are critical for the body to achieve energy homeostasis. Further, disruption of gut

nutrient sensing during high-fat feeding likely contributes to the development of obesity

and diabetes. Thus, it is crucial to further dissect the mechanisms through which

individual nutrients stimulate this gut-brain axis, and how they may contribute to the

pathology of metabolic diseases. To date, the most is known about intestinal lipid-

sensing mechanisms in the control of glucose homeostasis. Interestingly, however,

numerous studies have shown that intestinal protein is more potent than equicaloric

amounts of lipid or carbohydrate to stimulate gut peptide release155 and suppress

22

feeding91,156. In light of this, it is vital to further understand the role of dietary protein in

the gut-brain axis.

1.4 Intestinal protein sensing

High-protein diets increase circulating levels of both CCK and GLP-1157,158, and indeed,

luminal protein is a potent stimulus of the gut hormones CCK, GLP-1 and PYY158-161.

Intraduodenal infusion of protein hydrolysates or individual amino acids potently

suppresses feeding in rodents and humans91,92,161-163, the suppression of food intake by

intestinal protein requires CCK-dependent vagal afferent firing164, and high-protein diet

feeding is associated with activation of anorexigenic, CCK-responsive NTS neurons165.

However, the reduction of food intake by high-protein feeding is also associated with a

rise in GLP-1, and the potential role of GLP-1 is less clear158,166.

At the level of the intestinal epithelium, various sensory mechanisms have been

proposed to link luminal amino acids and proteins to the release of gut peptides. Gut

mucosal cells express a number of GPCRs sensitive to amino acids, oligopeptides, or

both, as well as transporters for either free amino acids or oligopeptides. While many of

these protein sensors have been linked to gut peptide release in vitro, the relative

importance of each sensor in the physiological regulation of the gut-brain axis remains

unclear.

23

T1R1-T1R3, the α-gustucin-coupled “umami” receptor that senses amino acids in taste

buds of the tongue, is also expressed throughout the small intestine, specifically by

CCK-expressing EECs, and in vitro, blocking T1R1 blunts CCK secretion in response

to a number of aliphatic amino acids167,168. Individual amino acids are taken up by an

array of transporters with varying specificities, many of which remain to be identified.

However, one such transporter, B0AT1, has been cloned in both rodents and humans

and found to transport the bulk of neutral amino acids169. B0AT1 knockout mice have

decreased intestinal amino acid signaling and insulin release in response to feeding170,

and intestinal B0AT1 expression is altered in obesity171, suggesting that this transporter

could play a role in nutrient sensing, though more work is required to fully elucidate its

role. GPR93 is a GPCR expressed throughout the gut mucosal epithelium, and

specifically by CCK-expressing cells, which senses both individual amino acids and

small peptides and has been implicated in CCK release in vitro172,173. Likewise, CaSR

is a GPCR that was originally identified as a receptor for calcium, but that also senses

amino acids and small peptides174. CaSR is expressed by primary duodenal I-cells

isolated from mice, and mediates aromatic amino acid-induced CCK release56. CaSR

has also been shown to mediate peptide-induced CCK and GLP-1 release in vitro175-

177. While each of these protein sensors has been implicated in cell lines, future studies

are required to address their physiological role in gut peptide release in vivo.

A number of the aforementioned receptors sense both amino acids and oligopeptides,

and it remains unclear whether luminal proteins usually signal in the form of free amino

24

acids, small peptides, or both, to stimulate gut peptide release during digestion. An

intraluminal free amino acid solution stimulates vagal afferent firing in rodents178 and

induces gut peptide release in humans179, suggesting that individual amino acids can

induce gut-brain signaling. However, intraduodenal protein hydrolysate infusion also

stimulates CCK-dependent vagal afferent firing180,181, and one study showed that co-

infusion of the proteolytic enzyme trypsin decreased the CCK response to protein

hydrolysates by 60%182, suggesting that proteins signal at least partially in the form of

oligopeptides. Further, one in vitro study showed that removing the lower molecular

weight fractions from protein hydrolysates, which includes free amino acids, has no

effect on its ability to induce CCK release from STC-1 cells, and that a free amino acid

solution does not in fact induce CCK release from these cells176. Along with the fact

that non-metabolizable peptidomimetics (synthetic peptides) activate EECs and

stimulate CCK release183-185, these results suggest that luminal protein may in fact

signal mainly in the form of di- and tripeptides, prior to further lysis into free amino

acids, to initiate a gut-brain negative feedback axis.

Unlike amino acid transport, the uptake of di- and tripeptides in the small intestine is

solely mediated by one molecule, PepT1. In addition to mediating enterocyte peptide

absorption, PepT1 has been localized on CCK-secreting cells and is able to elicit

voltage-gated calcium entry in response to the uptake of peptides177. Indeed, non-

metabolizable PepT1 substrates stimulate CCK release from STC-1 cells in vitro58.

Further, PepT1 inhibition blocks firing of CCK-responsive vagal afferents in response to

25

intraduodenal protein hydrolysate infusion ex vivo181, further highlighting its likely role in

the initiation of gut-brain signaling in response to luminal protein.

26

Figure 1: Intestinal nutrient sensing in the regulation of metabolic homeostasis. Luminal nutrients signal via cell-surface receptors and transporters to stimulate the

release of gut peptide hormones from enteroendocrine cells in the gut mucosa, via

receptor-induced membrane depolarization, second-messenger signaling cascades or

intracellular nutrient metabolism. Once released, gut peptides act on receptors

expressed locally by vagal afferent fibers innervating the lamina propria to induce vagal

afferent firing. Vagal afferents terminate in the NTS of the hindbrain, where neuronal

signals are relayed via the hypothalamus and/or vagal efferents to suppress food intake

and hepatic glucose production to maintain energy homeostasis. High-fat diet feeding

disrupts this nutrient-signaling axis, most likely at the level of gut peptide receptor

signaling on vagal afferents. NTS, nucleus of the solitary tract; HYP, hypothalamus.

26

Chapter 2

Hypothesis and Aims

Upper small intestinal nutrient sensing mechanisms are implicated in the regulation of

food intake and whole-body glucose homeostasis, and are disrupted in high-fat diet

feeding, potentially contributing to the development of obesity and diabetes. More

specifically, upper intestinal lipids stimulate the release of CCK, which acts on CCK-1

receptors expressed by vagal afferent neurons to signal via a gut-brain axis to

suppress feeding and a gut-brain-liver axis to suppress hepatic glucose production.

Alternatively, little is known about the role of intestinal protein sensing mechanisms to

regulate metabolic homeostasis, despite the fact that acute dietary protein intake

improves glucose homeostasis in both rodents and humans. Given that upper intestinal

protein suppresses food intake via CCK-mediated vagal afferent firing, we propose that

while in the pre-absorptive state, dietary proteins trigger a neuronal network to regulate

whole-body glucose homeostasis.

General Hypothesis: Upper small intestinal protein sensing triggers a neuronal

network to lower hepatic glucose production and maintain glucose homeostasis in

healthy rodents in vivo (Figure 2).

27

Aims:

Aim 1: To evaluate whether upper small intestinal protein sensing regulates glucose

homeostasis.

Aim 2: To investigate whether upper small intestinal protein-sensing mechanisms

regulate glucose homeostasis through direct effects on glucose uptake or glucose

production.

Aim 3: To evaluate whether a neuronal network is required for upper small intestinal

protein sensing to regulate glucose homeostasis.

28

Figure 2: Schematic of the working hypothesis. Luminal protein is sensed by the

gastrointestinal tract prior to absorption into the portal circulation, signalling to activate

a neuronal network within the gut wall. This signal is then relayed via a gut-brain-liver

neuronal axis to directly suppress hepatic glucose production, improving whole-body

glucose tolerance. NTS, Nucleus of the solitary tract; NMDA; N-methyl D-aspartate;

DVC, dorsal vagal complex.

29

29

Chapter 3

Materials and Methods «

3.1 Animals

For all studies, male Sprague-Dawley rats were obtained from Charles River

Laboratories (Montreal, QC, Canada) at 8-weeks of age (250-270g). Rats were

individually housed and maintained for six days with a standard 12-hour light-dark cycle

and ad libitum access to water and rat chow (Harlan Teklad; % calories: 16% fat/60%

carbohydrate/24% protein; total caloric value: 3.1 kcal/g) in order to acclimatize to our

facilities prior to surgical manipulation. All animal protocols were reviewed and

approved by the Institutional Animal Care and Use Committee at the University Health

Network.

3.2 Surgical Procedure

3.2.1 Preparation of cannulae

Duodenal cannula: 18 cm of 0.04 inner-diameter/0.085 outer-diameter silicon tubing

(Sil-Tec) was overlapped with 1 cm of smaller, 0.025 inner diameter/0.037 outer-

diameter silicon tubing (Sil-Tec). A 1-cm2 piece of polypropylene mesh (Bard Davol)

30

30

was glued to the cannula at the intersection of the two tubing sizes, such that the small

tubing passed through a hole in the center of the mesh square. Vascular cannulae: 15

cm of PE50 polyethylene tubing (Clay Adams) was fitted with a 1.5 cm cuff extension of

15 mm inner-diameter Silastic silicon tubing (Dow Corning).

3.2.2 Cannulation surgery

Rats were anesthetized with an intraperitoneal injection of a cocktail of ketamine (60

mg/kg; Vetalar, Bioniche) and xylazine (8 mg/kg; Rompun, Bayer). The abdominal and

neck regions were shaved and sanitized with 70% ethanol and 10% providine-iodine

solution (Betadine). A 4 cm laparotomic incision was made along the ventral midline

through both the skin and abdominal muscle wall, exposing the GI tract within the

peritoneal cavity. The duodenum was isolated 6 cm distal to the pyloric sphincter. A

small hole was made in the intestinal wall using a 21-guage needle in an area with the

least vascularization to minimize bleeding, and the intestinal cannula was inserted into

this hole and anchored to the outer surface of the duodenum by applying tissue

adhesive (3M Vetbond) to the surrounding mesh. The duodenal cannula was flushed

with saline to ensure that the infusion flowed into the lumen of the duodenum with no

leakage. The duodenum was then repositioned within the peritoneal cavity and the

abdominal wall was closed with 4-0 silk sutures such that the cannula exited through

the abdominal wall between 2 continuous sutures. A 1.5 cm incision was made in the

back of the neck and the cannula was tunneled subcutaneously from the abdomen,

exiting the incision in the neck. The neck and abdominal incisions were closed with 4-0

silk sutures and the end of the cannula, which now exited through the back of the neck,

was closed with a metal pin.

31

A 3 cm incision was then made in the front of the neck, and through blunt dissection of

fat, connective tissue and muscle, the right jugular vein was isolated. The vessel was

ligated at the cranial end and loosely tied at the caudal end using 4-0 sutures, and the

ligatures were pulled taught to prevent blood flow in the isolated segment of the vessel.

A small incision was made in the vessel wall using microscissors, and a vascular

cannula filled with 1% heparinized saline was inserted into the vessel lumen until it

reached a position where blood freely flowed into the cannula when an attached

sampling syringe was pulled back. The cannula was then secured in place by

tightening the loosely tied ligature around the vessel and cannula. This procedure was

repeated with the right carotid artery. The two cannulae were then tunneled

subcutaneously with a 16-guage needle so that they exited the back of the neck on

either side of the intestinal cannula. The neck incision was closed with 4-0 sutures, and

the vascular cannulae were filled with 10% heparin solution to prevent clotting and

closed with metal pins. 10 mL of 0.9% saline was injected subcutaneously to help with

recovery and rats were returned to their cages.

Rats were given 4 days to recover from surgery with ad libitum access to water and

regular chow. The intestinal cannula was flushed with saline daily to prevent clogging.

Food intake and body weight were monitored daily and rats recovered to at least 90%

of their pre-surgical body weight prior to experiments.

32

3.3 Intestinal infusions and treatments

The following treatments were infused into the upper small intestine via the indwelling

cannula during the in vivo experiments, at a rate of 0.12 ml/min for 1 minute to fill the

dead space of the cannula followed by 0.01 ml/min to a total of 50 minutes: 1) 0.9%

saline, 2) 8% (w/v) casein hydrolysate 3) tetracaine (0.01 mg/mL) and 4) 8% (w/v)

casein hydrolysate + tetracaine (0.01 mg/mL). All of the treatments were adjusted to pH

5, which is the native pH of the rat upper small intestine186.

Casein was selected for its broad amino acid composition and because it is a common

dietary protein source. The hydrolyzed form of casein protein is composed of small

peptides and free amino acids and most closely resembles the partially digested form

in which dietary proteins reach the upper small intestine during digestion. The typical

amino acid analysis of casein hydrolysate (Sigma-Aldrich) can be seen in Table 1. The

dose of 8% (w/v) casein hydrolysate (0.32 kcal/mL) was selected because

intraduodenal 8% peptone stimulated vagal firing and suppressed food intake in

rodents in vivo181,187. 3.2 g of casein hydrolysate (Sigma-Aldrich) was solubilized in 40

mL of saline by lowering the pH to 1 through titration with 10 M HCl and subsequently

raising the pH to 5 through titration with 10 M NaOH.

The tetracaine dose was based on a previous study where 0.01 mg/min tetracaine

blocked intraduodenal lipid infusion to regulate glucose homeostasis138. Tetracaine

33

(Sigma-Aldrich) was dissolved in DMSO in a stock solution of 200 mg/mL, and 22.5 µL

of this stock solution was added to 3 mL of either saline or 8% casein hydrolysate.

3.4 Intravenous glucose tolerance test

Food was removed at 5 pm the night before the glucose tolerance test for an overnight

fast. Experiments took place at 10:00 AM. The experimental protocol can be seen in

Figure 3A. The intraintestinal infusion was initiated at t=-15 at a rate of 0.12 mL/min for

1 minute to fill the dead space of the intestinal cannula, followed thereafter by a rate of

0.01 mL/min, administered using a Harvard Apparatus PHD 2000 syringe pump. At t=0,

the glucose injection was administered (0.25g/kg, 20% glucose solution), over 10

seconds, via the jugular vein cannula. 0.4 mL blood samples were drawn from the

carotid artery cannula at t=-15, t=0 (prior to the glucose injection), t=2, t=5, and every 5

minutes thereafter until t=35, for plasma glucose and amino acid analysis (see below).

The experiment was terminated at t=35, or the end of 50 minutes of gut infusion. At the

end of the 50-minute experiment, rats were anesthetized via intravenous administration

of ketamine, and a 0.5 mL portal vein blood sample was drawn for plasma amino acid

analysis.

34

3.5 Pancreatic (basal insulin) euglycemic clamp

Rats were restricted to 15 g (~47 kcal) of food the night prior to the clamp experiment,

which took place at 9:00 AM. The clamp protocol can be seen in Figure 4A. An

intravenous infusion of [3-3H]-glucose (40 µCi bolus; 0.4 µCi/min; Perkin Elmer) was

initiated at t=0 and continued for the duration of the 200-minute experiment. From t=60

to t=90, once the [3-3H]-glucose has reached steady-state in the rat, 0.2 mL blood

samples were drawn every 10 minutes from the carotid artery for plasma glucose

analysis (see below) in order to obtain basal plasma glucose readings. At t=90, the

pancreatic clamp was initiated through continuous intravenous infusion of somatostatin

(3 µg/kg/min; Bachem) and insulin (1.2 mU/kg/min; porcine insulin; Sigma-Aldrich).

Somatostatin was infused to inhibit all endogenous insulin secretion while the

exogenous insulin infusion replaced insulin at basal levels. An intravenous infusion of

exogenous glucose (25% solution) was also initiated at t=90. Plasma glucose readings

were obtained every 10 minutes from t=90-200, and the exogenous glucose infusion

rate was adjusted as required to maintain euglycemia according to the basal glucose

readings. During the final 50 minutes of the experiment (t=150-200), the upper small

intestinal infusions were administered as previously described above. Additional 50 µL

plasma samples were obtained at all of the 10-minute time points for later plasma

specific activity analysis. At the end of the experiment, at time t=200, rats were

anesthetized via intravenous administration of ketamine, and a 0.5 mL portal vein blood

sample was drawn for plasma amino acid analysis. All infusions were administered

using a Harvard Apparatus PHD 2000 syringe pump.

35

3.6 Biochemical analysis

3.6.1 Plasma glucose

During in vivo experiments, blood samples were collected into heparinized Eppendorf

tubes and centrifuged (6000 rpm, 1 min) to separate the plasma. 10 µL of plasma was

then analyzed using a GM9 Glucose Analyzer (Analox Instruments), which utilizes the

glucose oxidase method, in order to obtain plasma glucose readings in situ.

3.6.2 Plasma amino acids

A protocol to measure plasma amino acid concentrations using the colorimetric

ninhydrin reaction was adapted from 188. Briefly, the assay is based on the principle that

ninhydrin reacts with equal affinity to the α-amino nitrogen groups of all amino acids,

with the exception of profile, producing a purple colour. This colour change can be

measured using a spectrophotometer and compared to known concentrations of α-

amino acids, resulting in a measure of total α-amino acids in the sample.

The specific protocol was as follows: The plasma samples were first deproteinized such

that the α-amine groups in the samples would be indicative of only free α-amino acids.

50 µL of plasma in 350 µL of ddH2O was deproteinized through the addition of 50 µL of

36

10% (w/v) sodium tungstate (Sigma-Aldrich) and 50 µL of 2N HCl, with successive

mixing. After standing for 5 minutes, samples were centrifuged (13000 rpm, 5 min,

4°C). 250 µL of supernatant was then combined with 125 µL of cyanide-acetate buffer

(pH 5.2) containing: 114.65 g sodium acetate (Sigma-Aldrich), 500 mL ddH2O, 36 mL

glacial acetic acid, 5g disodium EDTA (BioShop) and 20 mL cyanide solution (490 mg

sodium cyanide [Sigma-Aldrich] in 1 L ddH2O). 125 µL of ninhydrin reagent (3% (w/v)

ninhydrin [Sigma-Aldrich] in 2-methoxyethanol [Sigma-Aldrich]) was added. Tubes were

mixed and capped, and then heated in a boiling water bath for 15 minutes. Immediately

upon removal from the water bath, 3.75 mL cold diluent (50% isopropanol in ddH2O)

was added. Samples were mixed thoroughly by inversion, and transferred to 1 cm

cuvettes. In addition to the plasma samples, a “blank” sample was prepared as

described above but with 50 µL ddH2O replacing the 250 µL of deproteinized plasma,

and a “standard” sample was prepared using 50 µL of stock amino acid solution (26.8

mg glycine [BioShop] in 100 mL ddH2O) replacing the deproteinized plasma. All

cuvettes were read in a spectrophotometer (Genesys 10S VIS, Thermo Scientific) at

570 nm. Given that the standard sample contains a known concentration of 5 mg α-

nitrogen/100 mL, the following formula was then used to calculate the concentration of

α-nitrogen in the samples based on their absorbencies (E):

Plasma α-N (mg/100 mL) = (E test) – (E blank) X 5 (E standard) – (E blank)

37

3.6.3 Plasma [3-3H]-glucose specific activity

50 µL plasma samples were deproteinized through the addition of 100 µL Ba(OH)2 and

100 µL ZnSO4, vortexing, and centrifugation (13200 rpm, 5 min, 4°C). The supernatant

was then transferred to scintillation vials without lids and left for 48 hours to evaporate

under a fumehood. Evaporation rids the samples of water containing [3H] as a product

of glycolysis, such that only [3-3H]-glucose will contribute to the radioactivity in the

sample. 7.5 mL of Bio-Safe Scintillation Cocktail (Research Products International

Corp) was then added to the dry samples and the vials were counted in a LS6500

Multipurpose Scintillation Counter (Beckman Coulter).

3.7 Tracer-dilution calculations

The constant, radioactive [3-3H]-glucose tracer infusion reaches steady state in the

plasma and tissues of the rat within one hour. At this point, a steady state formula can

be applied to calculate glucose uptake and glucose production, whereby the rate of

glucose disappearance (Rd) is equal to the rate of endogenous glucose appearance

(Ra). The specific activity of the plasma samples collected during the basal period of

the clamp (t=60-90) can therefore be used to calculate glucose uptake and glucose

production at each time point as follows:

38

Ra = Rd = Tracer infusion rate (µCi/min) = Rates of Glucose production Specific activity (µCi/mg) and uptake (mg/kg/min)

During the clamp, when the exogenous glucose infusion is introduced, the rate of

glucose production must be calculated by subtracting the exogenous glucose infusion

rate (GIR), as follows:

Ra + GIR = Rd Ra = Rd – GIR = Rate of glucose production (mg/kg/min)

3.8 Statistical analysis

For the intravenous glucose tolerance test, the glucose concentration over time was

compared using a two-way anaylsis of variance (ANOVA) and was followed by a

Tukey’s post-hoc test when comparing more than two groups. The area under the

curve (AUC) was calculated with Prism software (GraphPad) as compared using an

unpaired Student’s t-test when comparing two groups or a one-way ANOVA when

comparing more than two groups. For the pancreatic clamp/tracer-dilution data

analysis, an average from t=60-90 was used as the “basal” values and an average from

t=180-200 was used as the “clamp” values. Comparisons between basal versus clamp

39

glucose uptake and glucose production were made using an unpaired Student’s t-test.

For plasma amino acid concentrations, the concentration at t=50 was compared

between saline versus casein hydrolysate treated rats using an unpaired Student’s t-

test. Prism software (GraphPad Software Inc.) was used for all statistical calculations.

A probability of P < 0.05 was accepted as significant. Values were presented as mean

± SEM.

40

Chapter 4

Results «

Upper small intestinal protein sensing improves glucose tolerance

To investigate the effects of upper small intestinal protein sensing mechanisms on

whole-body glucose homeostasis, we first developed a protocol incorporating a

simultaneous upper intestinal protein infusion and intravenous glucose tolerance test

(IVGTT) in healthy rodents in vivo. Sprague-Dawley rats were chosen as the rodent

model to allow for future comparisons between healthy and high-fat diet-fed rodents.

Upon high-fat feeding, Sprague-Dawley rats are susceptible to the development of

hyperphagia, insulin resistance and obesity, with less variability than other rat

strains189. Accordingly, only male rats were chosen as male Sprague-Dawley rats are

more susceptible than females to high-fat diet-induced hyperphagia189, and their

metabolism is not affected by variable sexual hormone cycles. All male Sprague

Dawley rats were obtained from Charles River Laboratories at 8 weeks of age, and

given exactly six days to acclimatize to our facilities prior to surgical manipulation. It is

now well understood that environmental history influences rodents’ metabolism and

their response to dietary challenges and should therefore be controlled190. On the

seventh day in our facilities, rats underwent jugular, carotid and small intestinal

cannulation surgeries. After 4 days

41

of recovery time during which they were fed an ad libitum regular chow diet, rats were

subjected to an overnight fast (food removed at 5 pm on day 4) followed by the gut-

infusion IVGTT experiment on the morning of day 5 (Figure 3A).

The IVGTT was selected to evaluate whole-body glucose homeostasis during an upper

small intestinal protein infusion, while circulating glucoregulatory hormones change at

will. The OGTT, while commonly utilized, is confounded by the rates of intestinal

glucose absorption and gastric emptying, thus producing more variable data than an

IVGTT191, which evaluates the effects of a treatment specifically on the tolerance of

circulating glucose, controlled only by glucose uptake and glucose production.

Additionally, the IVGTT provides the possibility of future mathematical modeling of our

results using the “minimal model” system developed by Bergman and colleagues to

evaluate the relative roles of insulin-dependent and insulin–independent effects7.

Finally, and most importantly, intravenous administration of glucose bypasses the

upper small intestine and does not interfere with our intestinal treatments. Rats were

fully awake throughout the IVGTT experiments, as general anesthetics can have both

direct and indirect effects on metabolic studies192. The IVGTT protocol is summarized

in Figure 3A: Briefly, the intravenous glucose injection was administered after 15

minutes of a 50-minute intestinal infusion of the given treatment, and blood samples

taken frequently to assess plasma glucose levels. The upper intestinal treatments were

administered directly into the lumen via the small intestinal cannula, first at a rate of

0.12 mL/min for one minute to fill the dead space of the cannula with the treatment and

then at a rate of 0.01 mL/min for a total of 50 minutes. Infusion of 8% casein

42

hydrolysate for 50 minutes at this rate did not increase portal or systemic free amino

acids (Figure 3B). This is consistent with previous findings that 20% intralipid

administered into the duodenum at the same rate does not increase portal or systemic

free fatty acids138. Thus, we successfully developed an in vivo model to evaluate the

effects of pre-absorptive protein-signaling mechanisms in the upper small intestine on

the regulation of whole-body glucose homeostasis.

As assessed by the IVGTT, upper small intestinal infusion of casein hydrolysate

significantly improved glucose tolerance compared to the control group receiving

intraintestinal saline, where plasma glucose levels were significantly lower from times

10-25 (Figure 4A). Further, the integrated area under the curve (AUC) of the glucose

collapsed over time was significantly reduced in protein-treated rats compared to saline

(Figure 4B). These results were independent of any differences in body weight or

cumulative post-surgical food intake (Table 2). Given that the protein treatment did not

raise circulating amino acids, our data demonstrate that upper small intestinal protein

sensing regulates whole body glucose homeostasis to improve glucose tolerance.

43

Upper small intestinal protein sensing suppresses glucose production

An improvement in glucose tolerance during an IVGTT can be accounted for entirely by

an increase in peripheral glucose uptake and/or a suppression of endogenous glucose

production. Furthermore, the effects of intestinal nutrient sensing on either glucose

uptake or glucose production may be either dependent on, or independent of, an

incretin effect to increase insulin secretion. Thus, our lab has developed a protocol

utilizing a pancreatic basal-insulin euglycemic clamp technique in conjunction with

tracer-dilution methodology to assess the direct effects of intestinal nutrient infusions

on whole-body glucose kinetics (glucose uptake and glucose production)138 (Figure

5A). To evaluate the mechanisms through which intestinal protein improves glucose

tolerance, we administered the 50-minute upper intestinal casein hydrolysate treatment

in a clamp setting, in a new group of rats that underwent the same handling, surgery

and recovery protocol as those receiving the IVGTT. The clamp experiment took place

the morning of day 5 post-surgery, and rats were restricted to 15 g (~47 kcal) of food

the night before to ensure a similar post-absorptive status. The 200-minute clamp

protocol is summarized in Figure 5A: Briefly, the [3-3H]-glucose is infused and allowed

to reach steady state, somatostatin is infused to inhibit endogenous insulin secretion

and insulin is replaced to basal levels. In the final 50 minutes of the clamp, the 50-

minute gut infusion is administered at the same rate as during the IVGTT, and an

exogenous glucose infusion is administered as required to maintain euglycemia. At

basal circulating free amino acid and insulin levels (Figure 3B), upper small intestinal

infusion of casein hydrolysate increased the exogenous glucose infusion rate (GIR)

44

required to maintained euglycemia 5-fold compared to a control saline infusion (Figure

5B). Analysis of the tracer-dilution data revealed that this increase in GIR was entirely

accounted for by a 48% suppression of hepatic glucose production (Figure 6A,B), as

glucose uptake was unaltered (Figure 6C). There were no differences in body weight or

food intake between groups (Table 2). Thus, upper small intestinal protein sensing

regulates glucose homeostasis through directly suppressing hepatic glucose

production, independent of changes in circulating insulin.

Upper small intestinal protein sensing requires local, neuronal signaling to

regulate glucose homeostasis

Given that upper intestinal lipids suppress glucose production via a gut-brain-liver

neuronal axis138, we next evaluated the role of a neuronal axis to mediate the effects of

protein sensing on whole-body glucose homeostasis. Gut-peptides released in

response to luminal nutrients act on locally expressed receptors to stimulate vagal

afferent neuron firing9, and co-infusion of tetracaine, a topical anesthetic, with

intraintestinal nutrients negates the ability of nutrient sensing to suppress food intake

and lower glucose production by blocking afferent neuronal signaling to the brain138,193.

We therefore co-infused tetracaine with upper small intestinal casein hydrolysate during

the IVGTT in a new group of rats, to assess whether a gut-brain neuronal axis is

required for intestinal protein sensing to improve glucose tolerance. Infusion of

tetracaine alone (1 mg/mL, 0.01 mg/min) had no effect on glucose tolerance compared

45

to saline, while co-infusion of tetracaine with 8% casein hydrolysate reversed the

improvement in glucose tolerance by casein hydrolysate (Figure 7A). The integrated

AUC was reversed to that of the saline-treated rats (Figure 7B). Therefore, intestinal

protein activates local, neuronal signaling in order to regulate glucose homeostasis and

improve glucose tolerance, further supporting that the signaling mechanism of protein

sensing is localized within the gut.

46

A B

Figure 3: Gut infusion-IVGTT protocol to evaluate the effects of upper small intestinal protein sensing on glucose tolerance. A, Experimental protocol for the

intravenous glucose tolerance test with upper intestinal infusions. IVGTT, intravenous

glucose tolerance test; SI, small intestine. B, Systemic and portal vein plasma free

amino acid levels in response to the 50-minute upper intestinal saline (n=10) or casein

hydrolysate (n=10) infusion. Values expressed as mean ± SEM. No differences among

groups as determined by unpaired Student’s t-test.

47

A B

Figure 4: Upper small intestinal infusion of protein improves glucose tolerance. A, Plasma glucose concentrations over time during the intravenous glucose tolerance

test in rats receiving an upper small intestinal infusion of saline (n=10) or casein

hydrolysate (n=10). B, The integrated area under the curve (AUC). Values expressed

as mean ± SEM. * p < 0.05, ** p < 0.005 compared to saline as determined by two-way

ANOVA.

48

A B

Figure 5. Upper small intestinal casein hydrolysate increases the glucose infusion rate during the basal insulin euglycemic clamp. A, Experimental protocol

for the basal insulin euglycemic clamp. B, The glucose infusion rate required to

maintain euglycemia during the clamp with upper small intestinal infusion of saline

(n=7) or casein hydrolysate (n=7). Values expressed as mean ± SEM. * P < 0.05

compared to saline as determined by unpaired Student’s t-test.

49

A B

C

m

Figure 6. Upper small intestinal casein hydrolysate suppresses hepatic glucose production. A, B, C, Rate of glucose production (A), suppression of glucose

production expressed as the percent decrease from basal (B) and clamp rate of

glucose uptake (C) with upper small intestinal infusion of saline (n=7) or casein

hydrolysate (n=7). Values expressed as mean ± SEM. * P < 0.05 compared to saline as

determined by unpaired Student’s t-test.

50

A

B

Figure 7: Upper small intestinal protein requires local, neuronal signaling to regulate glucose tolerance. A, Plasma glucose concentrations over time during the

intravenous glucose tolerance test with upper small intestinal infusion of saline (n=10),

casein hydrolysate (n=10), tetracaine (n=8) or casein hydrolysate with tetracaine (n=8).

B, The integrated area under the curve (AUC). Values expressed as mean ± SEM. * P

< 0.05 compared to saline, # P < 0.05 compared to tetracaine, t P < 0.05 compared to

51

casein hydrolysate + tetracaine, as determined by two-way ANOVA and Tukey post

hoc test.

Table 1: Typical amino acid content of Casein enzymatic hydrolysate from Sigma-Aldrich (Product 22090).

Amino acid Typical concentration (%w/w)

Glutamic acid 16.5

Proline 8.5

Leucine 6.5

Lysine 6.4

Aspartic acid 5.5

Valine 5.3

Isoleucine 4.5

Serine 4.5

Phenylalanine 3.8

Threonine 3.6

Arginine 2.9

Alanine 2.4

Methionine 2.4

Histidine 2.1

Tyrosine 1.8

Glycine 1.6

Tryptophan 0.95

Cystine 0.67

52

Table 2: Body weight and cumulative post-surgical food intake of the groups of rats that underwent either the intravenous glucose tolerance test or clamp protocol.

  SAL  (IVGTT)  

CAS  (IVGTT)  

TET  (IVGTT)  

CAS  +  TET  

(IVGTT)  

SAL  (clamp)  

CAS  (clamp)  

Body  weight  (g)   279  ±  6.0   280  ±  3.9   274  ±  4.9   263  ±  7.9   295  ±  6.7   298  ±  10.4  

Food  intake  (g)   26.1  ±  3.5   23.3  ±  3.2   21  ±  2.9   18  ±  3   30.0  ±  2.9   29.2  ±  2.6  

Values expressed as mean ± SEM. Saline (SAL). Casein hydrolysate (CAS).

Tetracaine (TET). Intravenous glucose tolerance test (IVGTT). Basal insulin euglycemic

clamp (clamp).

53

Table 3: Plasma glucose concentrations during the basal insulin euglycemic clamp.

Basal Saline Casein hydrolysate

Glucose (mg/dL) 142.4 ± 6.5 137.8 ± 6.3

Clamp Saline Casein hydrolysate

Glucose (mg/dL) 159.3 ± 5.5 147.2 ± 2.8

Values expressed as mean ± SEM.

54

Chapter 5

Discussion

To date, the role of dietary protein in the regulation of glucose homeostasis remains

unclear. Although there is evidence that the resulting increase in plasma amino acids

may contribute to gluconeogenesis and induce insulin resistance27,194, many studies

show that high-protein diet feeding in fact lowers fasting glucose and HbA1c levels and

improves glucose tolerance28,29,32-34. The mechanisms of this improvement in glucose

control are unknown, but may involve acute negative feedback by intestinal protein

sensing. Indeed, using a model of healthy, male, Sprague-Dawley rats, we here

demonstrate that upper small intestinal protein sensing regulates whole-body glucose

homeostasis. Specifically, a 50-minute upper small intestinal infusion of casein

hydrolysate improved glucose tolerance as assessed by an IVGTT in comparison to

rats given a control infusion of saline, independent of any differences in food intake or

body weight.

Analysis using the ninhydrin method indicated that this improvement in glucose

tolerance occurred independent of a rise in either portal vein or systemic free amino

acid levels. Circulating amino acids can act on other targets to regulate glucose

homeostasis, such as in the hypothalamus to lower hepatic glucose production via a

brain-liver neuronal network195. In addition, vagal afferents innervating the portal vein

respond to portal amino acid infusion196,197, implying the presence of portal amino acid

55

sensors. However, the fact that amino acid levels were not raised by the 50-minute

casein hydrolysate infusion suggests that protein acts via gut-localized sensing

mechanisms to regulate whole-body glucose homeostasis during the IVGTT.

It is, however, important to note that the plasma samples were deproteinized, and the

ninhydrin assay measured only free amino acids, not accounting for those that make up

oligopeptides. It can therefore not be ruled out that di- and tripeptides absorbed by the

gut epithlium intact are reaching the portal vein within 50 minutes. During digestion of a

mixed meal, 90% of oligopeptides absorbed by epithelial cells intact are hydrolyzed

intracellularly and reach the portal vein as free amino acids198. However, given that

apical peptide transporters are upregulated by the presence of luminal oligopeptides199,

our intraintestinal casein hydrolysate infusion may lead to increased flux of peptide

absorption, causing release of excess peptides into the circulation, which are missed

by the ninhydrin assay, and which can act on peripheral targets to regulate glucose

homeostasis. Therefore, future studies should employ a control group where the 8%

casein hydrolysate treatment is infused at the same rate directly into the portal vein

(assuming 100% leakage), to rule out post-absorptive effects improving glucose

tolerance. Nevertheless, the present results suggest that dietary proteins act locally

within the upper small intestine to improve whole-body glucose tolerance in healthy

rodents.

56

These findings are in line with previous studies showing that acute protein intake can

improve glucose tolerance. Consumption of a protein hydrolysate beverage lowers the

glucose response to a subsequent mixed meal43,200, and the addition of protein

hydrolysates to a standard carbohydrate beverage lowers the glucose response42.

Furthermore, consumption of a 5g high-protein diet meal results in a lower glucose

response than a 5g low-protein meal as early as 30 minutes after the initiation of

feeding45. One proposed mechanism of this lowered glucose response is a protein-

induced lowering of gastric emptying rate201, which would then slow the appearance of

glucose into the circulation. However, our current findings indicate that when the

confounding variable of gastric emptying is removed by intravenous injection of the

glucose bolus, intestinal protein signals to improve whole-body glucose tolerance, and

gut protein sensing may therefore contribute to the improvement in glucose response

seen in these acute feeding studies.

The improvement in intravenous glucose tolerance induced by small intestinal protein

sensing must be accounted for by a suppression of endogenous glucose production

and/or increase in peripheral glucose uptake. There are multiple mechanisms through

which gut protein sensing may alter either of these factors. The improvement in glucose

turnover may be secondary to a change in circulating glucoregulatory hormones, given

that protein intake stimulates the release of insulin202, and may do so via pre-absorptive

gut peptide-mediated signaling. GLP-1, which is released in response to luminal

protein, stimulates the release of insulin either via direct action on β-cells or a local

neuronal axis mediated by hepatoportal vagal afferents and pancreatic efferents203, in

57

order to improve glucose tolerance129. Additionally, protein-induced CCK release may

contribute to this incretin effect given that CCK is implicated in stimulating postprandial

insulin secretion both in vitro and in vivo134,204. In many instances, however, acute

protein intake improves glucose homeostasis independently of a rise in insulin

levels43,45. Indeed, protein-induced gut peptide release could regulate glucose

homeostasis independently of insulin, such as via a GLP-1-mediated gut-brain-muscle

neuronal axis to increase muscle uptake and metabolism130,205. However, the most like

likely explanation may be a suppression of hepatic glucose production rather than an

increase in glucose uptake, given that both fatty acid and glucose sensing within the

small intestine suppresses hepatic glucose production via a gut-brain-liver neuronal

axis, independently of any effects on glucose uptake, during a basal insulin pancreatic

clamp13,138.

To investigate whether upper small intestinal protein sensing regulates either glucose

uptake or glucose production, independently of any changes in insulin, the 50-minute

casein hydrolysate infusion was next given during a basal-insulin euglycemic clamp.

During the clamp, upper small intestinal protein infusion increased the glucose infusion

rate required to maintain euglycemic, and this was completely accounted for by a

suppression of hepatic glucose production, and glucose uptake remained unchanged.

Thus, upper small intestinal protein-sensing mechanisms directly regulate hepatic

glucose production, and this likely contributes to the improvement in whole-body

glucose tolerance in response to intestinal protein during the IVGTT. However, the

contribution of this suppression of hepatic glucose production to the improvement in

58

glucose tolerance seen during the IVGTT remains unknown. In order to address the

contribution of hepatic glucose metabolism in future experiments, metabolic pathways

regulating glucose production in the liver could be blocked during the IVGTT.

Upper small intestinal lipids initiate a gut-brain-liver neuronal axis, mediated by vagal

afferents, NMDA receptor transmission in the DVC and the hepatic vagal efferent to

suppress hepatic glucose production138. Thus, a similar neuronal axis may mediate

intestinal protein to suppress glucose production. Indeed, co-infusion of the local

anesthetic tetracaine completely reversed the improvement in glucose tolerance by

upper intestinal protein infusion, indicating that pre-absorptive protein initiates a

neuronal network at the level of the gut to exert its effects on glucose homeostasis.

Alternatively, upper small intestinal infusion of tetracaine alone did not affect glucose

homeostasis during the IVGTT, indicating that the contribution of basal vagal afferent

firing to glucoregulatory control is negligible, whereas a postprandial increase in firing is

required for nutrients to regulate glucose homeostasis. These findings are consistent

with a recent study investigating the role of gut neuronal signaling in humans. When

implanted in the stomach, the electrical DIAMOND® device senses incoming food and

responds with electrical stimulation of the gut wall. After 5 weeks, this postprandial

neuronal stimulation resulted in an improvement in glycemic control (lower HbA1c) in

type 2 diabetic individuals206. Thus, the present findings are likely to be conserved in

humans and may be important in the context of disease.

59

Future studies are required to dissect the involvement of a gut-brain or gut-brain-liver

neuronal axis in the regulation of glucose tolerance and glucose production. The ability

of upper small intestinal lipids to suppress hepatic glucose production is attenuated not

only by co-infusion of tetracaine but also by blocking NMDA receptor transmission

within the NTS of the hindbrain or by surgical ablation of the hepatic vagal efferent,

implicating a gut-brain-liver axis mediated by vagal afferents, NMDA transmission in the

NTS and the hepatic vagus138. Pharmacological inhibition of NTS NMDA receptors, by

NTS infusion of the noncompetitive high-affinity antagonist MK-801, and hepatic

vagotomy surgery should thus be utilized during future studies to investigate whether a

similar gut-brain-liver axis mediates intestinal protein to suppress hepatic glucose

production during the pancreatic clamp.

The precise mechanisms through which upper small intestinal casein hydrolysate

initiates a neuronal network regulating glucose homeostasis remain unknown. Casein

hydrolysate is a partially hydrolyzed form of protein, containing both individual amino

acids and oligopeptides, for both of which there are sensory machineries expressed by

GI epithelial cells that are implicated in gut peptide release. A number of amino acid-

specific GPCRs are expressed by EECs and mediate CCK release in response to

amino acids in vitro56,168,172,173, which vary in their affinity for different classes of amino

acids. Casein hydrolysate is composed of a broad range of amino acids, with the

highest proportion being Glutamic acid followed by the aliphatic amino acids Proline,

Leucine and Lysine. Indeed, the gut epithelium expresses glutamate receptors, which

mediate gut peptide release and vagal afferent firing in response to luminal glutamate

60

infusion and possibly to high protein meals207, one of which is T1R1-T1R3, the gut-

expressed α–gustducin coupled GPCR and “umami” taste receptor168. The role of

aliphatic amino acids such as Leucine in the gut-brain axis is less clear given that

although intraduodenal infusion of protein hydrolysate suppresses feeding, infusion of

Leucine, Arginine or Alanine independently does not suppress feeding162. Alternatively,

infusion of the aromatic amino acids Phenylalanine or Tryptophan alone suppresses

feeding162. Indeed, Phenylalanine and Tryptophan stimulate CCK release and suppress

feeding via a vagal afferent-dependent neural axis in both rodents and humans161,208,

and Phenylalanine infusion is in fact more potent than casein hydrolysate to suppress

feeding in rodents92. These findings indicate that although Tryptophan and

Phenylalanine are in lower proportion than other amino acids in casein hydrolysate,

they may be the signaling molecules through which this protein solution initiates a gut-

brain axis. Future studies should look at whether infusion of either of aromatic amino

acids alone can equally or more potently regulate glucose tolerance and glucose

production in comparison to the casein treatment. In vitro, aromatic amino acids are

sensed by the GPCR CaSR, which is expressed by EECs, in order to stimulate CCK

release56, thus the role of this transporter in protein-induced glucoregulation should

also be investigated. Alternatively, uptake and metabolism of amino acids by gut

epithelial cells may be required, as in the case of intestinal lipid sensing, where

metabolism to LCFA-CoA is required for lipids to stimulate CCK release and lower

glucose production138. Indeed, hypothalamic leucine infusion lowers plasma glucose via

a suppression of hepatic glucose production, and this is dependent upon metabolism to

the intermediate α-ketoisocaproic acid and subsequently to acetyl-coA and malonyl-

CoA195, which is known to increase cytoplasmic accumulation of LCFA-CoA, increasing

61

flux into the lipid sensing pathway to suppress glucose production. Given that upper

intestinal accumulation of LCFA-CoA activates a CCK-dependent gut-brain-liver axis to

suppress glucose production138, intestinal amino acids may act through a similar

pathway to regulate glucose homeostasis via metabolism to malonyl-CoA.

However, protein hydrolysates or solutions of small peptides are more efficiently

absorbed than amino acid mixtures, suggesting that a large amount of dietary protein is

absorbed in the form of oligopeptides rather than hydrolyzed into amino acids within the

lumen. Therefore, luminal protein hydrolysate may additionally or alternatively signal in

the form of oligopeptides. Indeed, co-infusion of the proteolytic enzyme trypsin

attenuates the CCK response to luminal protein hydrolysate182, and non-metabolizable

oligopeptides stimulate CCK release183. The protein-sensor PepT1 is likely involved in

sensing these oligopeptides, given that this di- and tripeptide-specific transporter

mediates CCK release as well as vagal afferent firing in response to intestinal protein

hydrolysate58,181. In order to evaluate the role of PepT1, future studies will employ co-

infusion of the competitive non-translocated inhibitor 4-aminomethylbenzoic acid (4-

AMBA) with upper small intestinal protein during both the IVGTT and clamp protocols.

To further confirm the role of PepT1, the PepT1 protein will be knocked down in the

upper small intestine with a lentiviral shRNA infection during the gut cannulation

surgery as confirmed in Côté et al209. Finally, upper small intestinal infusion with non-

metabolizable substrates of PepT1 such as the cephalosporin antibiotic Cefaclor or the

synthetic dipeptide Gly-Sar during the IVGTT or clamp would confirm whether substrate

binding and activation of PepT1 per se initiates the neuronal network to regulate

62

glucose homeostasis or whether subsequent intracellular metabolism of peptides is

required.

Identification of the precise mechanisms through which protein initiates this negative

feedback pathway on glucose metabolism is important to potentially unveil therapeutic

targets at the level of the gut to lower hyperglycemia in disease. Indeed, this gut-brain-

liver axis has recently been demonstrated to have important therapeutic relevance.

Both metformin and resveratrol activate gut-localized signaling pathways to lower

hepatic glucose production via a neuronal gut-brain-liver axis in high-fat diet-fed and

type 2 diabetic rodents10,209. Furthermore, in humans, 12-week treatment with delayed-

release Metformin, which targets the lower bowel where little absorption occurs, is

more effective in lowering glucose levels in type 2 diabetic patients than 12 weeks of

commercially available immediate-release or extended-release Metformin11. These

results confirm those in rodents suggesting that the primary effect of Metformin on

glucose homeostasis may be via gut-localized signaling. These exciting new studies

confirm that gut nutrient signaling pathways may have pharmacological relevance in

disease and that it is urgent to elucidate the precise molecular mechanisms through

which intestinal nutrients signal, and how signaling defects in high-fat feeding may be

bypassed by selectively targeting these pathways.

While we have confirmed that upper small intestinal protein sensing regulates

intravenous glucose tolerance and suppresses glucose production during the

63

pancreatic clamp, future studies are required to determine the physiological relevance

of protein sensing in the context of high-protein feeding. As previously mentioned, re-

feeding of a high-protein diet following an overnight fast lowers the glucose response in

healthy male Sprague-Dawley rats in comparison to low-protein feeding45.

Interestingly, blocking hindbrain glucagon receptor signaling partially attenuates this

improvement in glucose tolerance in the later time points (60 minutes after re-

feeding)45, but has no effect on the earlier time points. Thus, the same fasting re-

feeding protocol should be employed in future studies to investigate the role of

intestinal protein sensing in mediating this early improvement in glucose control. Once

we have identified the specific mucosal sensing mechanisms that mediate protein to

improve glucose homeostasis, using the techniques described above, this pathway

should be blocked during re-feeding of a high-protein diet through pharmacological

inhibition or viral knockdown. To investigate how intestinal protein sensing might

contribute to the glucoregulatory capacity of longer-term dietary interventions, knockout

mouse models could be employed. For example, PepT1 mice that lack the transporter

in the intestines and kidney have been developed, and do not display any obvious

abnormalities in size, weight fertility or viability210. It would be interesting to investigate

whether the lack of intestinal PepT1 might become important in the context of high

protein diet feeding, whereby PepT1 knockout attenuates the ability of long-term high

protein diet intervention to improve glucose regulation.

In summary, the present study shows for the first time to our knowledge that upper

small intestinal protein initiates a neuronal network to improve glucose tolerance and

64

suppress hepatic glucose production, likely via pre-absorptive signaling to initiate gut-

brain and/or gut-brain liver signaling axes. These findings demonstrate that the GI tract

and brain have parallel protein-sensing mechanisms to lower hepatic glucose

production given that central amino acid infusion lowers glucose production during the

same basal insulin pancreatic clamp protocol195,211. While blocking neuronal signaling

with tetracaine completely reversed the effects of intraintestinal protein infusion on

glucose tolerance in the context of a dose which does not rise portal or systemic

plasma amino acids, it is important to note that under physiological high-protein feeding

conditions, where amino acids rise in the circulation, other protein-sensing mechanisms

such as in the brain likely become important.

It will be of great interest in future studies to test the efficacy of upper small intestinal

protein infusion to regulate glucose homeostasis in high-fat diet-fed rats. 3 days of

high-fat diet feeding reverses the effects of upper small intestinal lipid infusion to lower

glucose production, by disrupting the lipid-sensing pathway at the level of intestinal

CCK-1 receptor signaling138,143. Interestingly, however, high protein diet feeding

effectively improves glucose homeostasis in diet-induced obese rodents212,213, and

acute protein intake lowers blood glucose levels in type 2 diabetic humans214,215,216.

Thus, we propose that unlike lipid sensing, intestinal protein sensing mechanisms may

in fact remain intact in rodent models of metabolic disease, suggesting that protein-

sensing pathways may differ from lipid sensing, and further justifying the importance to

investigate the potential therapeutic relevance.

65

Chapter 6

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