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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
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.
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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
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
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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.
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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
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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
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intraduodenal protein hydrolysate infusion ex vivo181, further highlighting its likely role in
the initiation of gut-brain signaling in response to luminal protein.
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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.
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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).
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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.
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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.
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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)
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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.
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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.
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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%
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:
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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
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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.
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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
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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
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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.
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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)
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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
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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.
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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.
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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.
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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.
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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.
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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
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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
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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.
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