Published January 28, 2015
DIGESTIVE PHYSIOLOGY OF THE PIG SYMPOSIUM:
Secretion of gastrointestinal hormones and eating control1
R. E. Steinert,*2,3 C. Feinle-Bisset,*2 N. Geary,† and C. Beglinger‡2
*University of Adelaide Discipline of Medicine and Centre of Clinical Research Excellence in Nutritional Physiology,
Interventions and Outcomes, Adelaide, SA 5005, Australia; †Zielackerstrasse 10, 8603 Schwerzenbach, Switzerland; and
‡Department of Biomedicine and Division of Gastroenterology, University Hospital Basel, Basel, 4030, Switzerland
ABSTRACT: Nutrient ingestion triggers numerous
changes in gastrointestinal (GI) peptide hormone
secretion that affect appetite and eating. Evidence for
these effects comes from research in laboratory animals,
healthy humans, and, increasingly, obese patients after
Roux-en-Y gastric bypass (RYGB) surgery, which has
marked effects on GI hormone function and is currently
the most effective therapy for morbid obesity. Increases in
cholecystokinin (CCK), glucagon-like peptide-1 (GLP1), and peptide tyrosine tyrosine (PYY) and decreases
in ghrelin secretion after meals are triggered by changes
in the nutrient content of the intestine. One apparent
physiological function of each is to initiate a reflex-
like feedback control of eating. Here we briefly review
this function, with an emphasis on the controls of their
secretion. Each is secreted from enteroendocrine cells
that are directly or indirectly affected by caloric load,
macronutrient composition, and other characteristics of
ingested food such as fatty acid chain length. In addition,
digestive hydrolysis is a critical mechanism that controls
their secretion. Although there are relatively few data
in agricultural animals, the generally consistent results
across widely divergent mammals suggests that most of
the processes described are also likely to be relevant to
GI hormone functions and eating in agricultural animals.
Key words: cholecystokinin, ghrelin, gastrointestinal
peptides, glucagon-like peptide-1, peptide tyrosine tyrosine
© 2013 American Society of Animal Science. All rights reserved.
INTRODUCTION
Among the many functions of gastrointestinal
(GI) hormones that are secreted in response to food
ingestion is that of providing feedback signals to the
brain that control eating, in particular, the reflexive
control of meal onset and termination. Many recent
advances in understanding the physiology of these
1Based on a presentation at the preconference symposium titled
“Gut Chemosensing: Integrating nutrition, gut function and metabolism
in pigs” preceding the 12th International Symposium on Digestive
Physiology of Pigs in Keystone, Colorado, May 29–June 1, 2012, with
publication sponsored by Lucta S.A. (Barcelona, Spain), the American
Society of Animal Science, and the Journal of Animal Science.
2R.E. Steinert is supported by a Novartis Grant, C. Feinle-Bisset
by a National Health and Medical Research Council of Australia
(NHMRC) Senior Research Fellowship (grant no. 627002), and
C. Beglinger by grant no. 320030-132960/1 of the Swiss National
Science Foundation.
3Corresponding author: [email protected]
Received October 22, 2012.
Accepted December 20, 2012.
J. Anim. Sci. 2013.91:1963–1973
doi:10.2527/jas2012-6022
hormones have come from studies in normal-weight
humans and in obese subjects who have undergone
Roux-en-Y gastric bypass (RYGB) surgery, which is
currently the most effective therapy for morbid obesity
(Colquitt et al., 2005; Pournaras and Le Roux, 2009).
One of the mechanisms thought to contribute to the
beneficial effects of RYGB surgery is earlier exposure
of the mid jejunum and more distal small intestine
to nutrient chyme and, as a consequence, substantial
changes in the secretory patterns of GI hormones,
including ghrelin, cholecystokinin (CCK), glucagonlike peptide-1 (GLP-1), and peptide tyrosine tyrosine
(PYY; Pournaras and Le Roux, 2009; Peterli et al.,
2012). This review provides a brief overview of the
roles of these hormones in the control of eating, with
an emphasis on the mechanisms that control their
secretion. Although most of the data stem from studies
in humans and laboratory rodents, the consistent
results across widely divergent mammals, including
the modest literature in pigs, cattle, and sheep, indicate
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Steinert et al.
that most of the processes described here are also likely
to be relevant to agricultural animals.
GHRELIN
Ghrelin is secreted mainly by X/A-like cells in the
fundus of the stomach and, unique among GI hormones,
stimulates eating. Its concentrations increase during
fasting and decrease after meals, and intravenous
infusion of ghrelin in both humans and animals
significantly increases meal size, consistent with an
acute role in meal initiation (Tschop et al., 2001; Ueno
et al., 2005; Kirchner et al., 2012). In addition, basal
ghrelin concentrations are decreased with overfeeding
and increased by BW loss, indicating that ghrelin may be
a tonic signal linked to body adiposity (Cummings et al.,
2002; Williams et al., 2006). In 2008, Yang et al. (2008)
showed that the enzyme ghrelin O-acetyltransferase
(GOAT) catalyzes the conversion of ghrelin into its
biologically active acylated form, indicating that GOAT
is important in the pathogenesis of obesity. However, the
physiological relevance of the effect of ghrelin on eating
requires more research. For example, it is still unknown
whether the physiological preprandial rise in circulating
ghrelin is sufficient to trigger eating and whether ghrelin
acts peripherally or centrally to control eating (Date et
al., 2002; Arnold et al., 2006; Langhans and Geary, 2010;
Kirchner et al., 2012)
Although the stomach is the major site of ghrelin
secretion, it does not appear to contain the sensing
mechanisms that suppress ghrelin secretion after meals,
because nutrient exposure of the stomach alone is
not sufficient to suppress ghrelin (Shiiya et al., 2002;
Williams et al., 2003; Overduin et al., 2005). Rather, the
available data indicate that the signals mediating the
postprandial suppression of ghrelin originate further
distally in the small intestine (Williams et al., 2003;
Feinle-Bisset et al., 2005; Cukier et al., 2008; Ryan et al.,
2012). In addition, changes in plasma insulin, intestinal
osmolarity, or enteric neural signaling may mediate the
suppression of ghrelin (Gelling et al., 2004; Cummings,
2006). A study by Shiiya et al. (2002) indicated that
chemosensory mechanisms that respond to plasma
glucose concentrations control ghrelin secretion.
Cummings and coworkers (Callahan et al., 2004)
studied healthy volunteers receiving liquid meals with
widely varied caloric content but equivalent volume
and found that postprandial ghrelin suppression was
proportional to the ingested caloric load. In addition, in
studies in humans and rats they showed that carbohydrate
and protein suppress ghrelin concentrations more
effectively than lipid (Monteleone et al., 2003; Overduin
et al., 2005; Cummings, 2006). In our own recent
experiments, we found that intraduodenal lipid infusions
are associated with a marked and progressive suppression
of plasma ghrelin. Moreover, ghrelin suppression depends
on lipid digestion into FFA and FFA with a chain length
≥C12 are more potent than FFA with a chain length <C12
(Feltrin et al., 2006; Degen et al., 2007).
Available data indicate that plasma ghrelin
concentrations in cattle are similarly affected by fasting
and energy repletion (Wertz-Lutz et al., 2006, 2008,
2010). For example, a study in dairy cows revealed
a linear decline in pasture intake and postprandial
ghrelin concentrations in cows fed nutrient-concentrate
supplements (Roche et al., 2007). In addition, Wertz-Lutz
et al. (2006) reported that intravenous infusion of ghrelin
that produced fasting concentrations of plasma ghrelin
were sufficient to increase time spent eating in cattle.
Finally, a polymorphism in a gene controlling ghrelin
deacylation was reported to be associated with average
energy intake and weight gain in cattle (Lindholm-Perry et
al., 2012). The effects of fasting and refeeding on ghrelin
secretion and the eating-stimulatory effect of ghrelin
administration have been replicated in pigs (Zhang et
al., 2007; Dong et al., 2009). Fasting and refeeding also
affect ghrelin secretion in sheep, but tests of the effects
of ghrelin administration on eating have produced both
positive (Sugino et al., 2004; Grouselle et al., 2008) and
negative (Iqbal et al., 2006; Krueger and Melendez, 2012)
results, so that further work on this issue is warranted.
CHOLECYSTOKININ
Intestinal CCK is considered the paradigmatic GI
control of eating because of the extensive evidence
for its role in satiation in animals and humans (Gibbs
et al., 1973; Kissileff et al., 1981; Geary, 2004).
Cholecystokinin was purified as a 33 AA peptide by Mutt
and Jorpes (1968). Although many different forms of
CCK, including CCK-8, CCK-22, CCK-33, and CCK58, were subsequently found in either the GI tract or in
the brain, where it acts as a neurotransmitter (Rehfeld et
al., 2007), recent data indicate that CCK-58 may be the
only relevant endocrine form, with all the smaller forms
of CCK being proteolytic fragments (Eysselein et al.,
1990; Liddle, 1997; Stengel et al., 2009). Intestinal CCK
is released from duodenal I-cells. In addition to eating
control, the putative physiological functions of CCK
include the stimulation of the exocrine pancreas, liver,
and gallbladder to secrete digestive enzymes and bile
acids as well as the control of gut motility and gastric
emptying (Liddle et al., 1986; Beglinger, 1994).
In 1973, Gibbs et al. (1973) first implicated CCK
in the control of eating by showing that intraperitoneal
injections of CCK into rats immediately before meals
reduced meal size in a dose-dependent manner without
signs of aversions and without reducing water intake in
Gastrointestinal hormones and eating
thirsty rats. Since then, the ability of CCK signaling via
the CCK-1 receptor to inhibit eating has been studied
extensively in many species, with most of these studies
using CCK-8, which is a full agonist for the CCK-1
receptor. Importantly, administration of CCK-1 receptor
antagonists before meals selectively increases meal size
in rats, mice, and humans. In addition, in both humans
and rats, abnormalities of the CCK-1 receptor gene are
associated with increased meal size, increased food
intake, and overweightness in humans and rats (Miller
et al., 1995; Moran et al., 1998; Marchal-Victorion et al.,
2002; de Krom et al., 2007). However, CCK-1 receptor
knockout mice are not hyperphagic or obese (Kopin et
al., 1999). Whether this is related to the development
of compensatory mechanisms is not clear. The satiating
effect of exogenous CCK is very short-lived. The halflife of CCK is only about 2 to 3 min, and food intake
is not affected when it is injected >15 min before a
meal. Furthermore, the effect does not carry over from
one meal to the next (Cummings and Overduin, 2007;
Woods and D’Alessio, 2008). Whether or not this would
limit the usefulness of CCK agonism for the long-term
control of food intake is uncertain.
Available data indicate that CCK acts via an
endocrine mode of action to inhibit eating in humans
(Beglinger and Degen, 2004; Geary, 2004) but mainly
via a paracrine mode in rats (Geary, 2004; Eisen et
al., 2005). Evidence for the latter conclusion includes
1) there are only few reports of increased plasma CCK
concentrations during meals in rats (Geary, 2004),
2) intravenous infusions of CCK-8 into the hepatic portal
vein failed to reduce food intake (Greenberg et al., 1987),
3) under conditions in which intravenous infusions of a
CCK-1 receptor antagonist failed to increase eating, local
administration of the CCK-1 receptor antagonist into
the superior pancreaticoduodenal artery increased food
intake (Cox, 1998), and 4) intravenous infusion of a CCK
antibody that decreased pancreatic amylase secretion
failed to increase food intake whereas infusion of a smallmolecule CCK-1 receptor antagonist under the same
conditions had both effects, which strongly indicates
a paracrine action because the antagonist but not the
antibody would escape the circulation (Reidelberger et
al., 1994). Cholecystokinin is thought to signal satiation
via vagal fibers that directly innervate the tissue in close
proximity to the site of CCK secretion. Vagal afferents
have been shown to express CCK-1 receptors, and
lesioning either the subdiaphragmatic vagal nerves or the
dorsal vagal complex in the hindbrain has been shown to
attenuate the anorectic effect of peripheral CCK (Smith
et al., 1981; Edwards et al., 1986). Cholecystokinin
may also limit food intake, in part, by slowing gastric
emptying, as observed in both rodents (Moran and
McHugh, 1982) and humans (Lal et al., 2004). Consistent
1965
with this hypothesis, Schwartz et al. (1993) showed
that intravenous CCK infusion and small gastric loads
synergistically increased vagal afferent firing in rats.
Liddle et al. (1985), who developed the first reliable
assay for plasma CCK, found that glucose, lipid,
protein, and AA all rapidly and potently stimulate CCK
release. Subsequent studies have confirmed this and
also demonstrated that protein and lipid are the most
potent secretogogues (Himeno et al., 1983; Hopman et
al., 1985; Pilichiewicz et al., 2007; Seimon et al., 2009).
In our own experiments in humans, we recently showed
that isoenergetic replacement of lipid by carbohydrate
in intraduodenal infusions increased plasma CCK
concentrations less than did lipid alone (Seimon et al.,
2009). The exact mechanisms underlying CCK secretion
are not fully understood. The majority of data indicate
that digestion of proteins to oligopeptides and AA is
required to efficiently release CCK (Gibbs and Smith,
1977; Cuber et al., 1990a; Darcel et al., 2005; Foltz et al.,
2008). Similarly, when fat hydrolysis was blocked via
lipase inhibition, CCK release was markedly suppressed
(Hildebrand et al., 1998; Matzinger et al., 2000; Feinle et
al., 2001, 2003). Moreover, fatty acids with chain length
of ≥12 carbons stimulated CCK release much more than
fatty acids with <12 carbon atoms (McLaughlin et al.,
1998, 1999; Matzinger et al., 2000; Feltrin et al., 2004).
The control of eating by CCK in pigs has been studied
extensively. As in humans, carbohydrates, proteins and
lipids all stimulate CCK secretion in pigs (Houpt, 1984;
Cuber et al., 1990b). The effects on eating of exogenous
CCK, CCK-1 receptor agonism and antagonism, and
intraduodenal fat infusion in pigs are all similar to their
effects in humans (Anika et al., 1981; Gregory et al.,
1989; Ebenezer et al., 1990; Parrott, 1993). In addition,
active immunization against CCK increased food intake
and body weight in pigs (Pekas and Trout, 1990). There
has been less work on the satiating effect of CCK in
ruminants. In cattle, fasting reduced and nutrient repletion
increased plasma CCK concentrations (Suominen et al.,
1998) and, under some conditions, changes in plasma
CCK concentrations were associated with changes
in food intake (Relling and Reynolds, 2007, 2008). In
several studies in sheep, peripheral CCK administration
did not reliably inhibit eating (Grovum, 1981; Baile and
Della-Fera, 1984; Relling et al., 2011), perhaps because
the ovine vagus appears to express predominately CCK2 rather than CCK-1 receptors (Farningham et al., 1993).
In both pigs (Parrott, 1994; Baldwin and Sukhchai, 1996)
and sheep (Della-Fera et al., 1981; Della-Fera and Baile,
1984), there is evidence that CCK acts centrally to inhibit
eating. In both species this appears to be a local effect of
neural CCK released in the brain rather than an endocrine
effect of intestinal CCK.
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GLUCAGON-LIKE PEPTIDE-1
Glucagon-like peptide-1 is synthesized and
secreted from endocrine L-cells, which are expressed
most densely in the distal ileum and colon (Drucker,
2006; Holst, 2007). The bioactive form of GLP-1 is
generated from GLP-1(1–37) and exists as 2 equipotent
circulating molecular forms, GLP-1(7–37) and GLP1(7–36NH2). Both forms are rapidly degraded by
dipeptidyl peptidase-IV (DPPIV) to GLP-1(9–37) or
GLP-1(9–36NH2). Dipeptidyl peptidase-IV is primarily
located on the luminal surface of vascular endothelial
cells and circulating in the plasma (Mentlein, 1999;
Drucker, 2006; Holst, 2007). In addition to its incretin
effect (Schmidt et al., 1985; Drucker, 2006; Holst, 2007),
GLP-1 has been shown to inhibit eating in mice and rats
(Turton et al., 1996; Meeran et al., 1999; Abbott et al.,
2005; Chelikani et al., 2005). In humans, an early report
by Flint et al. (1998) indicated that GLP-1 infusions
enhanced fullness and reduced energy intake during an
ad libitum test meal. Subsequent studies demonstrated
that GLP-1 suppressed energy intake in obese and type
II diabetic subjects (Gutzwiller et al., 1999; Naslund et
al., 1999). However, some studies failed to demonstrate
an anorectic effect of GLP-1 in humans (Naslund et al.,
1998; Long et al., 1999; Brennan et al., 2005). Studies
using specific GLP-1 antagonists are required to better
establish a physiological role for GLP-1 in eating control
in humans.
The anorectic effect of GLP-1 is thought to be
mediated by GLP-1 receptors (GLP-1R; Bullock et al.,
1996; Alvarez et al., 2005; Holst, 2007). The anorectic
effect of exogenous GLP-1 is absent in transgenic GLP1R deficient mice and can be abolished in intact mice
with exendin(9–39), a selective GLP-1R antagonist
(Baggio et al., 2004; Williams et al., 2009; Ruttimann et
al., 2010). In addition, exendin(9–39) alone stimulated
food intake in rats, at least under some conditions
(Williams et al., 2009; Ruttimann et al., 2010; Asarian
et al., 2012). Whether the crucial GLP-1R are in the
brain, on vagal afferents, or elsewhere is unknown.
As mentioned, DPPIV rapidly catabolizes GLP-1, and
its plasma half-life is only 1 to 2 min. It is, therefore,
unclear whether an endocrine mechanism of action via
the systemic circulation is possible or whether intestinal
GLP-1 exerts its effects via receptors before reaching the
liver. Indeed, there is accumulating evidence indicating
that endogenous GLP-1, similar to CCK, has a paracrine
effect on intestinal vagal afferents in rats (Rüttimann et
al., 2009; Punjabi et al., 2011).
The secretion of GLP-1 has been studied
extensively. It is released postprandially in response to
all 3 macronutrients (Feltrin et al., 2004; Pilichiewicz
et al., 2007; Ryan et al., 2012). Some studies indicate
that carbohydrate is the strongest GLP-1 secretogogue,
consistent with its role as an incretin (Elliott et al., 1993;
Herrmann et al., 1995; Brubaker and Anini, 2003), but
other studies report that lipid and protein also potently
stimulate GLP-1 (Feinle et al., 2002; Raben et al., 2003;
Feltrin et al., 2004; Blom et al., 2006; Lejeune et al.,
2006). As with ghrelin and CCK, lipid hydrolysis is a
critical step for GLP-1 secretion. Fatty acids of chain
length <12 carbons have little effect (Feltrin et al., 2004;
Beglinger et al., 2010).
Depending on the macronutrient, GLP-1 is secreted
in a biphasic pattern, with an early-phase secretion
within 15 to 30 min after meal onset and a more
prolonged second phase from 1 to 3 h postprandially
(Herrmann et al., 1995; Schirra et al., 1996; Brubaker
and Anini, 2003; Drucker, 2006; Holst, 2007). The
GLP-1 response to carbohydrate is rapid and short-lived
whereas the responses to lipid and protein are slower and
more sustained (Elliott et al., 1993; Schirra et al., 1996;
Brubaker and Anini, 2003; Feinle et al., 2003). Several
mechanisms have been suggested to control GLP-1
secretion. A neurohumoral “proximal-to-distal loop”
may account for the early-phase secretion (Schirra et al.,
1996; Brubaker and Anini, 2003) including activation
of proximal M1 muscarinic or nicotinic receptors or
proximal I-cells that release CCK to stimulate more
distal GLP-1 containing L-cells (Anini and Brubaker,
2003; Beglinger et al., 2010). Alternatively, Egan and
colleagues (Theodorakis et al., 2006) suggested that
after a high-carbohydrate meal, activation of the small
number of proximal L-cells may be sufficient to account
for the early peak of plasma GLP-1. This is supported
by human studies demonstrating that the time courses
of glucose-induced GLP-1 secretion and appearance
of glucose in the proximal small intestine are similar
(Schirra et al., 1996; Theodorakis et al., 2006). However,
one of our recent studies underlines the importance of
an extended length of small intestine exposed for GLP1 release (Little et al., 2006), so that during a 60-cm
proximal-segment infusion of glucose, plasma GLP-1
concentrations did not change from baseline whereas
GLP-1 concentrations increased when glucose was
given access to the entire small intestine. Alternatively,
the second-phase secretion may also depend on the
production of metabolites from unabsorbed nutrients
by the gut microflora, most importantly the short-chain
fatty acids, acetate, propionate, and butyrate (Delzenne
et al., 2010). Another recent hypothesis suggests that
bile acids mediate GLP-1 secretion. Parker et al. (2012)
and Katsuma et al. (2005) showed in vitro that bile acids
induce an increase in GLP-1 by a TGR5 dependent
pathway. In addition, Adrian et al. (2010) found that
intrarectal infusion of taurocholic acid increased plasma
GLP-1 concentrations in obese and type 2 diabetic
Gastrointestinal hormones and eating
subjects. Others suggested that increases in plasma bile
acids may contribute to weight loss and the improvement
in glycemic control after RYGB surgery (Nakatani et al.,
2009; Patti et al., 2009; Pournaras et al., 2012).
The physiology of GLP-1 secretion has been studied
extensively in pigs and appears very similar to that in
humans (Holst, 2007). In a widely cited study in pigs,
Hansen et al. (1999) estimated that only 25% of the
secreted amount reaches the portal circulation and that
40 to 50% of that is destroyed in the liver, so that only
10 to 15% enters the systemic circulation. The GLP-1
agonist liraglutide inhibited eating in obese Göttingen
mini-pigs (Raun et al., 2007). We know of no reports
that GLP-1 or treatment with a GLP-1 agonist inhibits
eating in ruminants although in one study in dairy cows,
abomasal infusions of soybean oil led to increased
plasma GLP-1 concentrations and decreased DM intake
(Relling and Reynolds, 2008), and in another study in
sheep, there was a tendency for a reduction in DM intake
during 7-d infusions of GLP-1 (Relling et al., 2011).
PEPTIDE TYROSINE TYROSINE
Peptide tyrosine tyrosine was first isolated from
porcine intestine in 1982 by Tatemoto and named
“peptide tyrosine tyrosine” for the tyrosines at each end
(Tatemoto, 1982). Peptide tyrosine tyrosine is secreted
from endocrine L-cells of the distal ileum and colon,
most of which co-express GLP-1 (Adrian et al., 1985,
1987). In humans, about 60% of circulating PYY is
PYY(3–36), which results from cleavage of PYY(1–36)
by DPPIV (Medeiros and Turner, 1994; Ballantyne,
2006). Peptide tyrosine tyrosine (3–36) is inactivated by
nonspecific peptidases (Medeiros and Turner, 1994) and
has a half-life in plasma of approximately 9 min (Adrian
et al., 1985; Ballantyne, 2006).
In 2002, Batterham et al. (2002) first showed that
peripheral infusions of PYY(3–36) that generated
postprandial plasma concentrations reduced food intake
in humans. They also demonstrated that peripheral and
central injections of PYY(3–36) inhibited food intake
in rats and mice. Although some researchers could
initially not reproduce the eating-inhibitory effect in
rodents (Tschop et al., 2004), the anorectic action of
PYY(3–36) has now been shown in rats, mice, and other
species (Abbott et al., 2005; le Roux et al., 2006; Vrang
et al., 2006; Karra et al., 2009). However, it remains
unclear whether the anorectic effect of PYY represents
a pharmacological or a physiological phenomenon
in humans. We evaluated the effect on eating after
intravenous infusion of graded doses of PYY(3–36)
designed to mimic postprandial PYY secretion (Degen
et al., 2005). The results clearly indicated that significant
reductions in food intake were present only at plasma
1967
concentrations above those after a high-calorie meal.
Moreover, nausea was a common side effect, especially
at larger doses, indicating that at least some of the effects
of PYY on appetite might be due to malaise. Therefore,
whether or not PYY has a physiological role in eating
control in humans remains to be determined, and studies
using specific PYY receptor antagonists will be required
to fully characterize its physiological role in eating.
Peptide tyrosine tyrosine (1–36) is thought to exert
its effects via at least 3 different neuropeptide Y (NPY)family receptors (Y1, Y2, and Y5) whereas PYY(3–36)
has been shown to be selective for only one, Y2 (Dumont
et al., 1995; Ballantyne, 2006). This Y-receptor subtype
selectivity is thought to explain the different effects of
PYY(1–36) and PYY(3–36) on eating; that is, PYY(1–
36) increases eating by binding to orexigenic Y1 and Y5
receptor subtypes whereas PYY(3–36) decreases eating
via anorexigenic Y2 receptors (Cummings and Overduin,
2007). Several data are consistent with this hypothesis.
For example, Scott et al. (2005) found that Y2 receptor
antagonism completely blocked the eating-inhibitory
effect of PYY(3–36), and Batterham et al. (2002) showed
that PYY(3–36) inhibited eating in wild-type mice but
not in Y2-receptor knockout mice. The anorectic action
of PYY(3–36) may be mediated by vagal afferents.
Receptors for Y2 have been shown to be expressed on
vagal fibers, and vagotomy attenuated the anorectic effect
of peripheral PYY(3–36; Abbott et al., 2005; Koda et al.,
2005). Like GLP-1, PYY may also affect eating via direct
effects on gut motility (Witte et al., 2009).
Adrian et al. (1985) first showed in healthy humans
that test meals of increasing caloric content caused a
dose-related increase in plasma PYY. They also found
that lipids caused the greatest increase, protein caused
a more moderate increase, and glucose solution caused
only a transient and minor release, which has been
replicated by several groups (MacIntosh et al., 1999;
Essah et al., 2007; Brennan et al., 2012). As with
ghrelin, CCK, and GLP-1, PYY secretion depends on
lipid hydrolysis and fatty acid chain length ≥12 carbons
(Feltrin et al., 2006; Degen et al., 2007).
Plasma concentrations of total PYY or PYY(3–36)
increase to an initial peak 15 to 30 min after test meals,
reach a second peak approximately 60 to 90 min after
meals, and remain increased for up to 6 h (Adrian et
al., 1985; Degen et al., 2005; Ballantyne, 2006). As
with GLP-1, the secretion of PYY may be stimulated
indirectly via duodenally activated neuroendocrine
mechanisms or via bile acids as well as by direct contact
of nutrients with L-cells (Lin et al., 2000; Ballantyne,
2006). Although the patterns of secretion of PYY and
GLP-1 are similar, as expected because both peptides
are produced and secreted by L-cells, they are not
identical under all conditions. As mentioned previously,
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Steinert et al.
carbohydrates strongly trigger GLP-1 secretion but
are weak stimuli for PYY release (Adrian et al., 1985;
Herrmann et al., 1995; Brubaker and Anini, 2003),
indicating that subpopulations of L-cells may exist that
respond differently to a given nutrient stimuli. Indeed,
Eissele et al. (1992) found that GLP-1 and PYY were coexpressed in only some enteroendocrine cells in rat, pigs,
and humans. Another important consideration in this
context is that the patterns of secretion of GLP-1 and
PYY do not translate into the same patterns of plasma
concentrations of their bioactive forms because DPPIV
inactivates GLP-1 but activates PYY (Ballantyne, 2006).
SUMMARY AND CONCLUSIONS
It is increasingly recognized that nutrient stimuli
in the GI tract trigger a wide range of changes in GI
peptide hormone functions that influence appetite and
eating and that these effects are of clinical significance.
In particular, the mechanisms controlling the secretion
of gut hormones including ghrelin, CCK, GLP-1, and
PYY are now thought to be of importance for the success
of RYGB surgery in reversing obesity and metabolic
disease. In recent years, the use of direct nutrient
infusions into different areas of the GI tract and other
methods have provided new insights concerning the
roles of individual nutrients and specific GI segments in
the control of release of these peptides. For example, as
reviewed herein, carbohydrates, proteins, and lipids have
different potencies as ghrelin, CCK, GLP-1, and PYY
secretagogues, and lipid digestion and fatty acid chain
length importantly affect their secretagogue potencies.
However, it is important to emphasize that elucidating
the physiological control of GI peptide secretion is only
part of the challenge of understanding the roles of these
molecules in eating. As mentioned previously, paracrine
as well as endocrine signaling plays an important role
in eating control (Geary, 2004; Punjabi et al., 2011). In
addition, which aspect of the secretory patterns (e.g., rate
of change, specific time window) is critical for effective
signal transduction is unknown.
Functional interactions certainly also have a role as,
for example, has been convincingly demonstrated for the
synergistic effect of gastric fill and CCK on stimulation
of vagal afferent activity (Moran and McHugh, 1988;
Schwartz et al., 1993). These factors presumably
account for the frequent failure to detect correlations
between GI peptide concentrations and the amount eaten
during single meals (Seimon et al., 2010; Lemmens et
al., 2011). In addition, most of the work we reviewed
does not consider the effects of these peptides on food
reward. It is increasingly recognized that GI nutrient
sensing provides positive feedback signals that condition
food preferences and stimulate eating (Berthoud, 2011;
Sclafani and Ackroff, 2012). Moreover, GI peptides may
affect food reward via receptors outside the gut. For
example, endocrine GLP-1 may affect GLP-1 receptors
in the taste buds to influence the perception of sweet
taste (Martin et al., 2009), and a variety of evidence
links central ghrelin signaling to food reward (Egecioglu
et al., 2010). Emerging data from RYGB surgery also
indicate that alterations in GI nutrient sensing produced
by the surgery can change the rewarding effects of foods
in both rats (Shin et al., 2011) and humans (Miras et al.,
2012; Ochner et al., 2012). Finally, these pleiotropic
hormones have a variety of other effects as well, such
as actions on gastric emptying or glucose metabolism
(Ballantyne, 2006; Cummings and Overduin, 2007;
Holst, 2007), that may indirectly influence the more
direct effect of the hormone on eating.
Our review predominately concerned laboratory
rodents and humans. The extent to which these data are
relevant to animal husbandry is unclear. The several
consistent results across widely divergent mammals
indicate that most of the GI mechanisms involved in the
control of eating in humans and laboratory rodents are
likely to be relevant in agricultural animals. Nevertheless,
there are differences (e.g., the effects of central CCK
administration in sheep and pigs). Such differences
together with the general lack of knowledge concerning
the effects of GI peptides on eating in agricultural
animals indicate that this area urgently requires further
research. Such work seems likely to provide valuable
means to improve livestock health and productivity, for
example, by adjusting the nutrient composition of feed
in ways that optimize GI peptide hormone function.
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