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Neural substrates of anorexia
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Neural substrates of anorexia
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NOTE TO USERS
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unavailable from the author or university. The manuscript
was scanned as received.
95-104
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NEURAL SUBSTRATES OF ANOREXIA
by
Dawna Salter
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(NEUROSCIENCE)
August 2005
Copyright 2005 Dawna Salter
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UMI Number: 3196887
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Table of Contents
List of Figures and Tables iii
Abbreviations vi
Abstract ix
Chapter One: General Introduction 1
Chapter Two: Dehydration Anorexia and Glucoregulation
Introduction 30
Materials and Methods 33
Results 40
Discussion 51
Chapter Three: Differential Anatomical Patterns of Neuronal
Activation After 2-Deoxy-D-Glucose in Anorexic Animals
Introduction 60
Materials and Methods 62
Results 67
Discussion 74
Chapter Four: Attenuation of Feeding Responses after
Hypothalamic Injection of Neuropeptide Y is Site-Specific
in Anorexia
Introduction 85
Materials and Methods 87
Results 94
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Discussion 103
Chapter Five: Orexigenic Effects of Central AgRP Injections
Are Reversed By Anorexia
Introduction 112
Materials and Methods 114
Results 120
Discussion 130
Chapter Six: General Conclusions 138
Bibliography 145
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List of Figures and Tables
Figure 2.1 Timeline for experiments on EU and DE animals given 2-
deoxyglucose..................................................................................................... 39
Figure 2.2 Hypertonic saline ingestion inhibits feeding in response to
overnight starvation.......................................................................................... 42
Table 2.1 The effects of drinking 2.55 saline on body weights and mean
nocturnal food intake..................................................................................... 43
Figure 2.3 The feeding response to various doses of 2 deoxyglucose 44
Figure 2.4 The feeding response to doses of 2 deoxyglucose injections
repeated at 7 day intervals............................................................................ 45
Figure 2.5 The effects of 5 days of hypertonic saline ingestions on the
feeding response to 2 deoxyglucose injections........................................... 47
Table 2.2 The effects of drinking 2.55 saline on plasma glucose and
corticosterone responses to injections of vehicle or 2 deoxyglucose 49
Figure 2.6 The incremental increase in mean plasma glucose
concentrations above pre-injection values.................................................. 51
Figure 2.7 The incremental increase in mean plasma corticosterone
concentrations above pre-injection values..................................................... 52
Figure 3.1 Fos immunoreactivity after 2DG administration in euhydrated and
dehydrated animals......................................................................................... 68
Figure 3.2 Fos immunoreactivity after 2DG administration in euhydrated and
dehydrated animals.......................................................................................... 69
Figure 3.3 Mean number of Fos positive neurons after vehicle or 2DG
administration in euhydrated or dehydrated animals................................... 72
Figure 3.4 Mean number of Fos positive neurons after vehicle or 2DG
administration in euhydrated or dehydrated animals.................................... 73
IV
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Figure 4.1 Timeline for NPY experiments 91
Figure 4.2 Mapping of NPY injections sites.............................................. 93
Figure 4.3 The mean cumulative food intake in EU and DE animals four
hours after 0.5 ^ig NPY or vehicle injections targeted to the PVH 95
Figure 4.4 The mean cumulative food intake in EU and DE animals four
hours after 1.0 p,g NPY or vehicle injections targeted to the PVH.............. 96
Figure 4.5 The mean cumulative food intake in EU and DE animals four
hours after 0.5 p,g NPY or vehicle injections targeted to the LHApf 97
Figure 4.6 The mean cumulative food intake in EU and DE animals four
hours after 1.0 pg NPY or vehicle injections targeted to the LHApf 98
Figure 4.7 Correlation in DE animals between the development of
spontaneous anorexia and the inhibition of the feeding response after 1.0 pg
NPY injection to the LHApf.............................................................................. 99
Figure 4.8 Mean latency after injection of NPY until the initiation of eating in
EU and DE animals......................................................................................... 101
Figure 4.9 Mean cumulative food intake one hour after water is returned to
DE animals after vehicle injection or two doses of NPY directed towards
PVH or LHApf................................................................................................. 102
Figure 5.1 Timeline for fluid and food intake experiments for animals given a
3v injection of vehicle or one of three doses of A gR P ..................................118
Figure 5.2 NPY and AgRP mRNA hybridization in the arcuate nucleus of the
hypothalamus................................................................................................ 123
Figure 5.3 Hypertonic saline ingestion inhibits feeding 3 hours after 3v
AgRP injections........................................................................................... 124
Figure 5.4 DE inhibits feeding in response to 3v AgRP injection 125
Figure 5.5 Dose-responsive changes in body weight after 3v injection of
AgRP are different in EU and DE animals................................................... 126
v
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Figure 5.6 A high dose of AgRP stimulates fluid intake in EU, but not DE
animals, three hours following a single 3v injection................................... 128
Figure 5.7 72 hour fluid intake after different dose AgRP injection in EU and
DE rats.............................................................................................................. 129
Figure 5.8 Food intake 1 hour after water is returned to DE animals 130
Figure 6.1 Circuit diagram illustrating neural substrates involved in refeeding
after a period of negative energy balance.................................................... 139
Figure 6.2 Circuit diagram illustrating neural substrates involved in refeeding
after a period of negative energy balance continue to stimulate increased
production and release of NPY in the hypothalamus.................................. 141
Figure 6.3 Circuit diagram illustrating neural substrates involved in refeeding
after a period of negative energy balance................................................... 144
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Abbreviations
a-MSH a-melanocyte-stmulating hormone
2DG 2 deoxy-D-glucose
aCSF Artificial cerebral spinal fluid
AgRP Agouti related peptide
AP Area Postrema
ARH Arcuate nucleus of the hypothalamus
AVP Vasopressin
BNST Bed nucleus of the stria terminalis
BNSTfus Bed nucleus of the stria terminalis, fusiform part
CART Cocaine and amphetamine regulated transcript
CNTF Ciliary neurotrophin factor
CORT Corticosterone
CRH Corticotropin releasing hormone
CSF Cerebral spinal fluid
DE Dehydration
DMX Dorsal motor nucleus of the vagus
D-SAP Anti-dopamine beta-hydroxylase
E Epinephrine
EU Euhydrated
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FR Food restricted
GC Glucocorticoids
FIB Hindbrain
HIV Fluman immunodeficiency virus
IL Intraleukin
IR Immunoreactivity
LFIA Lateral hypothalamic area
LFIApf Lateral hypothalamic area, perifornical area
MC % Melanocortin 3 and 4 receptor
MCH Melanin-concentrating hormone
MnPO Median preoptic nucleus
NaCI Sodium chloride
NE Norepinephrine
NPY Neuropeptide Y
NT Neurotensin
NTS Nucleus of the solitary tract
O/FI Orexin/hypocretin
OT Oxytocin
OVLT Organum vasculosum of the lamina terminalis
PB Parabrachial area
PBN Parabrachial nucleus
POMC Pro-opiomelanocortin derived peptide
PVFI Paraventricular nucleus of the hypothalamus
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PVHIp Paraventricular nucleus of the hypothalamus, lateral
parvicellular part
PVHmpd Paraventricular nucleus of the hypothalamus, medial
parvicellular part
PVHpml Paraventricular nucleus of the hypothalamus, posterior
magnocellular part
RM Repeated measures
SCP Superior cerebellar peduncle
SFO Subfornical organ
SOL Solitary tract
STAT Signal transducer and activator of transcription
TNF Tumor necrosis factor
VEH Vehicle
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Abstract
Anorexia is marked by a loss of appetite despite the presence of palatable
foods and a physiological need for energy replenishment. Rats that are
dehydrated by drinking 2.5% saline for up to 5 days progressively reduce
their spontaneous, nocturnal food intake despite having endocrine and
neuropeptidergic profiles that normally elicit food intake. In fact, euhydrated
control animals food-restricted to match the food eaten by DE animals show
identical profiles but demonstrate an appreciable appetite, while DE animals
remain anorexic until given back water to drink (Salter and Watts, 2003;
Watts et al., 1999). Therefore, we present a series of experiments using
DE-anorexia to study how mechanisms that normally activate compensatory
feeding behavior after a period of negative energy balance might be
inhibited. First, we demonstrate the neural circuits activated by DE
specifically inhibit feeding, but not other regulatory responses, following
metabolic challenge. Next, we examine neuronal activation patterns to
identify areas where activation is absent in DE. Our results suggest DE-
anorexia is not mediated by those leptin-sensitive neurons in the arcuate
nucleus of the hypothalamus, but rather those neurons the lateral
parvicellular paraventricular nucleus and the lateral hypothalamic area may
contribute to inhibitory feeding pathways in DE-anorexia. We then test the
integrity of the feeding response in DE animals through central injections of
x
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orexigenic agents. We show that the hypothalamus is differentially
desensitized to orexigens depending on the anatomical target and the extent
of anorexia. Finally, we show that DE-anorexic rats usually demonstrate a
robust feeding response within ten minutes after drinking water is replaced,
suggesting the compensatory eating seen after DE may be initiated by
mechanisms divergent from those hypothalamic networks that ordinarily
stimulate adaptive feeding responses after energy deficit.
In a broader sense, the experiments in this thesis are designed to address
the pathophysiology of neural circuits and transmitters in the disequilibrium
between energy intake and expenditure in anorexia. As such, we begin to
address fundamental questions as to how appetite can be initiated and
turned off, and thus inform study of both obesity and anorexia.
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CHAPTER ONE
GENERAL INTRODUCTION
It’s hardly news anymore that Americans are just too fat. If the
endless parade of articles, TV specials and fad diet books weren’t proof
enough or you missed the ominous warnings from the National Institutes of
Health, the Centers for Disease Control and Prevention and the American
Heart Association, a quick look around the mall, the beach or the crowd at
any baseball game will leave no room for doubt: our individual weight
problems have become a national crisis.
Michael D. Lemonick, Time Magazine, June 7, 2004
Today the prevalence of obesity is rising at a pace so accelerated, the
population’s loosing fight against excessive weight gain has now been
recognized by the World Health Organization as one of the top 10 global
health problems (Kelner, and Helmuth, 2003). Obesity dramatically increases
the risk for life-threatening diseases such as diabetes, heart attack, stroke
and some types of cancer (Hill, Wyatt, Reed, and Peters, 2003). The overÂ
consumption of calories, hyper-efficient storage of metabolic fuels and
decreased energy expenditure despite elevated body fat stores are hallmark
components of obesity.
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On the opposite side of the spectrum is the malnutrition that results from
prolonged anorexia. The term anorexia is often used synonymously with
anorexia nervosa, a condition of self-imposed starvation that occurs most
commonly in adolescent females. In reality however, the term anorexia
simply describes a lack of appetite despite a physiological need for energy
replenishment and the availability of palatable foods. The contribution of
anorexia to malnutrition complicates the prognosis of many clinical conditions
including HIV infection, cancer, end-stage renal disease, chronic pulmonary
disease, and chronic inflammatory bowel disease. Anorexia is a major
contributor to cachexia, a condition of advanced protein-calorie malnutrition
that leads to invasive, expensive nutrition support therapy, iatrogenic
infections, poor response to therapy and, if not reversed, results in mortality.
In fact, involuntary weight loss is an independently reliable prognostic factor
in patients with AIDS, obstructive pulmonary disease and cancer, particularly
gastric and lung (Herndon, Green, Chahinian, Corson, Suzuki, and
Vogelzang, 1998; Kotler, 2000; Persson, Johansson, Sjoden, and Glimelius,
2002; Plata-Salaman, 1996; Prescott, Almdal, Mikkelsen, Tofteng, Vestbo,
and Lange, 2002). Additionally, anorexia is increasingly being recognized as
a frequent contributor to poor health in geriatric medicine.
Obesity and anorexia can be viewed as two sides of the same coin, as both
are disorders of homeostatic energy regulation. In either case the same
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biological circuitry has gone awry. Normally, energy homeostasis is
maintained through enormously complicated biological processes designed
to regulate the energy intake achieved by eating food with maximal efficiency
in its use and allocation. Caloric intake and energy expenditure are not
usually in a fixed ratio, however as needs change with ongoing activities
while the available food supply is seldom composed of the same nutrient
quality or hedonistic value. Therefore, the systems designed to cope with a
continuously changing energy requirement and a heterogeneously composed
energy supply must match the energy intake and expenditure over both short
and relatively long periods of time. Energy that is used for activity, heat, and
metabolism is normally matched by food intake (Bray, and York, 1998;
Keesey, and Hirvonen, 1997). This implies the existence of a regulatory
system that responds both to internal homeostatic needs as well as external
environmental changes. The existence of such a regulatory system is
illustrated by the dynamic interplay between energy intake and its use, so
that energy balance is usually achieved with a degree of relative precision
(Rothwell, and Stock, 1979; Weigle, 1994). For example, an average-sized
young American male eats more or less a million calories a year and utilizes
roughly the same amount. In fact, if this same male gains or loses 2-3
pounds per year, the regulatory system designed to achieve energy
homeostasis has still achieved an error rate just slightly over 1 %. Under
normal circumstances a perturbation of energy balance results in altered
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neural networks that orchestrate compensatory mechanisms designed to
defend body weight in the form of food intake and metabolic rate alterations
(Bray et al., 1998; Keesey et al., 1997). However, consistent environmental,
genetic or hormonal factors can cause progressive derangement of central
signaling pathways, the result being homeostatic dysregulation. Obesity
results from high circulating adiposity signals failing to halt over-consumption
of foods and/or raise the resting energy expenditure. Likewise, in anorexia
the negative energy balance brought on by reduced food intake fails to
activate adaptive processes to initiate an increased drive to eat.
The central nervous system directs ingestive behaviors and does so by
integrating a variety of information from external and internal environments
with neuropeptidergic networks to coordinate energy homeostasis. In the
recent past there has been remarkable progress in understanding the
complex interplay of the numerous distributed neural circuits and the way
they operate to coordinate energy homeostasis. This includes identifying
which neural networks stimulate, inhibit, or switch eating behaviors and how
they operate in response to signals that link the needs for energy balance
with calorie intake. Neuroscientific research done over the past 20 years has
provided tremendous advances in our understanding of the underlying
organization of neural circuits that regulate food intake and how they can be
modulated by regulatory metabolic and sensory signals.
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NEURAL SUBSTRATES INVOLVED IN ENERGY BALANCE
At the most basic level, ingestion is a motivated behavior composed of motor
contractions of striate muscle that move the animal to actively seek food, use
stereotypic motor movements to lick, chew and swallow the food, and to
terminate the meal by moving to another behavior. However, along with
activity of spinal cord and cranial nerve nuclei motor neurons, are also the
activities of the two other motor system divisions, the autonomic motor
neurons that control smooth muscle, cardiac tissue and glandular secretion
and the neuroendocrine motor neurons responsible for pituitary release.
Each of these systems affect fundamental motor events underlying energy
balance (Swanson, 2000). Further, each motor system is modulated by
sensory information which is integrated within neuropetidergic circuits that
interconnect constituent cell groups distributed throughout the neuraxis.
These sets of networks link sensory information with substrates in the hind-
and mid-brain, the hypothalamus and telencephelon, and serve to enact
mechanisms to organize motor outputs to maintain and restore energy
homeostasis. It is also important to note that all sensory modulation occurs
within the framework of the animal’s behavioral state which anchors most
motor events within the limits of the light-dark schedule. The essential role of
the hypothalamus for receiving and processing modulatory neural inputs and
organizing the endocrine, autonomic and behavioral responses necessary for
energy balance has been accepted for years. In particular are three
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interconnected cell groups that have been implicated in the control of energy
homeostasis and eating behaviors- the paraventricular nucleus of the
hypothalamus (PVH ), lateral hypothalamic area (LHA) and the arcuate
nucleus of the hypothalamus (ARH).
Paraventricular nucleus
The PVH is a cell group positioned to regulate numerous aspects of energy
balance. A large proportion of corticotropin releasing hormone (CRH)
neuroendocrine motor neurons reside within the PVH that can stimulate the
secretion of glucocorticoids from the adrenal cortex (Watts, 1996), as well as
populations of thyrotropin-releasing hormone, oxytocin and somatostatin-
producing neurons, that participate in energy metabolism. The PVH also
houses pre-autonomic motor neurons with descending efferent projections to
the spinal cord and brain-stern to control many autonomic functions including
gut function (Swanson, 1987; Zhang, Fogel, and Renehan, 1999) . Finally,
the PVH is implicated in eating behavior. NPY or norepinephrine injected
into the PVH stimulates eating behavior (Leibowitz, Sladek, Spencer, and
Tempel, 1988; Stanley, and Leibowitz, 1984) and starvation or glucose deficit
increases NPY release in the PVH (Beck, Jhanwar-Uniyal, Burlet, Chapleur-
Chateau, Leibowitz, and Burlet, 1990; Kalra, and Horvath, 1998). Although it
remains unclear exactly how the PVH mediates eating, it most likely involves
descending projections to the mesencephalic reticular formation,
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periaquaductal gray, the parabrachial nucleus, hindbrain and spinal cord,
areas with locomotor pattern initiators and generators (Swanson, 1991;
Swanson, 2000). Likewise, the PVH contains sets of nuclei which receive
and process modulatory sensory-derived afferents and organize appropriate
motor output.
Lateral Hypothalamic Area
The LHA has been associated with the control of ingestive behaviors since
the early work of Anand and Brobek (Anand, and Brobeck, 1951) when it
was found that circumscribed lesions of the LHA resulted in decreased food
intake so severe, the animals would die of starvation. These results have
since been challenged by research suggesting that the catastrophic aphagia
following severe lesions was in fact due to the loss of fibers of passage
rather than the destruction of constituent neuronal populations within the LHA
(Bernardis, and Bellinger, 1996; Strieker, 1984). However, when more
cellularly-specific excitotoxic lesions are made in those areas of the LHA that
are most closely related to feeding, the resultant deficit is rather mild and the
animals can compensate with appropriate responses following food or water
deprivation (Winn, 1995). Interestingly however, these lesioned animals
can not mount appropriate compensatory responses to direct homeostatic
challenges that are signaled entirely through internal metabolic indicators,
such as those that accompany glucose utilization deficit or extracellular fluid
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depletion (Winn, 1995). This supports the idea that the LHA modulates
ingestive behaviors through its extensive anatomical connections throughout
the neuraxis, including monosynaptic projections to the several areas of the
cerebral cortex and strong projections to the periaquaductal gray,
parabrachial area, and the hindbrain (Saper, 1985; Swanson, 1987).
Included in these projections are those that arise from LHA neurons
producing melanin-concentrating hormone (MCH, (Bittencourt, Presse, Arias,
Peto, Vaughan, Nahon, Vale, and Sawchenko, 1992) and
orexin/hypocretin,(0/H,(de Lecea, Kilduff, Peyron, Gao, Foye, Danielson,
Fukuhara, Battenberg, Gautvik, Bartlett, Frankel, van den Pol, Bloom,
Gautvik, and Sutcliffe, 1998; Sakurai, Amemiya, Ishii, Matsuzaki, Chemelli,
Tanaka, Williams, Richarson, Kozlowski, Wilson, Arch, Buckingham, Haynes,
Carr, Annan, McNulty, Liu, Terrett, Elshourbagy, Bergsma, and Yanagisawa,
1998) two peptides that when injected into the brain, increase food intake
(Qu, Ludwig, Gammeltoft, Piper, Pelleymounter, Cullen, Mathes, Przypek,
Kanarek, and Maratos-Flier, 1996; Sakurai etal., 1998). The LHA also
houses several other types of neurons. Within the LHA are neurons
producing cocaine-and-amphetamine related transcript (CART) and
neurotensin (NT), two neuropeptides that will decrease food intake after
intracerebroventricular injections (Kristensen, Judge, Thim, Ribel,
Christjansen, Wulff, Clausen, Jensen, Madsen, Vrang, Larsen, and Hastrup,
1998; Morley, 1987) and neurons that increase levels of anorexigenic CRH
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and NT during the manifestation of dehydration-anorexia (Watts, Sanchez-
Watts, and Kelly, 1999). The fact that the LHA is composed of large and
heterogeneous populations of neurons has complicated efforts to define its
precise role in the control of ingestive behavior, however these observations
suggest the neurons of the LHA play an important role in integrating afferent
metabolic signals with higher-order mechanisms responsible for volitional
aspects of feeding, and that the diverse neuropeptidergic efferents of the
LHA influences both stimulatory and inhibitory networks underlying feeding
behaviors.
Arcuate nucleus
Copious amounts of research have shown the ARH is intimately involved in
the regulation of energy intake. Neurons residing in the ARH are direct
targets of peripheral signals of energy status and these neurons send this
afferent input to both the PVH and the LHA (Broberger, Johansen,
Johansson, Schalling, and Hokfelt, 1998b; ELIAS, Saper, Maratos-Flier,
Tritos, Lee, Kelly, Tatro, Hoffman, Ollmann, Barsh, Sakurai, Yanagisawa,
and Elmquist, 1998b). Several neuronal populations are housed in the ARH
that can affect homeostatic aspects of ingestive behaviors. Two of these
neuropeptides are synthesized in the same ARH neurons, NPY and the
endogenous melanocortin receptor 3 and 4 (MC3/4) antagonist Agouti-
related peptide (AgRP), and stimulate feeding behaviors. Two other
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neuropeptides, a-melanin stimulating hormone (a-MSH) and CART are
derivatives of Pro-opiomelanocortin (POMC) and are also co-expressed in
the same neurons (Elias, Lee, Kelly, Aschkenasi, Ahima, Couceyro, Kuhar,
Saper, and Elmquist, 1998a). a-MSH, CART and neurotensin are all
neuropeptides produced in the ARH that inhibit feeding behaviors. Based
on the heterogeneity of neuronal content and the fact that these neurons are
easily engaged by metabolic and hormonal signals of changing energy
status, the ARH is a component of both inhibitory and stimulatory networks
that are integrated into motor output for energy homeostasis.
Modulatory Interosensory Inputs
Long-term and short-term balance in energy intake, utilization and storage is
needed for survival. Energy intake should be balanced with output while
maintaining enough energy stores to survive if the next meal is not
immediately available. However, energy stores must also be regulated as
overabundance of fat stores increases morbidity and mortality in all species.
The physiological systems designed to regulate energy balance are
modulated dynamically by sensory input. Both short-term and long-term
internal need states are coded as peripheral signals and are transduced by
the central nervous system through neuropeptides and their receptors, and
are integrated into coordinated motor activity in neuroendocrine, autonomic,
and somatosensory systems.
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Viscerosensory: Visceral sensory information from the gut and liver is
relayed through ascending vagal fibers and processed in the nucleus of the
solitary tract (NTS) in the hindbrain. From there, visceral information is sent
to the PVH, either directly or through the hindbrain catecholaminergic
neurons or the parabrachial nucleus (Swanson, 1987) and PVH output is
modified accordingly. For example, stimulation of vagal fibers elicits a
neuroendocrine response in the PVH (Rinaman, Hoffman, Dohanics, Le,
Strieker, and Verbalis, 1995; Ueta, Kannan, Higuchi, Negoro, Yamaguchi,
and Yamashita, 2000) and produces satiety effects including inhibition of
gastric motility, reduced food intake and stimulation of pituitary secretion
(Reidelberger, Arnelo, Granqvist, and Permert, 2001). Vagal efferents also
provide information on peripheral fatty acid oxidation status which in turn
elicits changes in rates of feeding (Horn, Addis, and Friedman, 1999; Ritter,
Dinh, and Friedman, 1994; Tordoff, Rawson, and Friedman, 1991).
Glucose status: Although there are undoubtedly glucose receptors in the
periphery, a competitive glucose substrate block activates brainstem
catecholaminergic neurons (Ritter, Llewellyn-Smith, and Dinh, 1998). These
noradrenergic and adrenergic neurons send afferent information to the
hypothalamus (Sawchenko, and Swanson, 1982; Tucker, Saper, Ruggiero,
and Reis, 1987)and rapidly elicit autonomic, neuroednocrine and somatic
motor actions. Sympathoadrenal activation elevates circulating blood
1 1
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glucose, activation of the hypothalmo-pituitary axis stimulates glucocorticoid
secretion and animals seek food to eat after administration of 2-deoxy-D-
glucose (2DG), a glucose analogue that blocks glucose metabolism.
Together these motor activities serve to redistribute and replenish metabolic
supplies. Interestingly, lesions of the PVH or the ARH do not effect eating
behaviors after 2DG administration (Bugarith, Dinh, Li, Speth, and Ritter,
2005; Calingasan, and Ritter, 1992) and evidence suggests the PVH may not
be a critical component for the autonomic motor events after this block of
glucose utilization (DiRocco, and Grill, 1979; Ritter, Bugarith, and Dinh,
2001a). Because the PVH houses NE motor neurons, however, suggests
the PVH is critical for the endocrine response to glucoprivation. A constant
supply of glucose to the brain is mandatory for survival, and the only way to
restore glucose stores is through food intake, so it likely other cell groups
participate in organizing this important behavior, possibly the LHA.
Fluctuations in circulating glucose levels are directly sensed by neurons in
ventromedial hypothalamus, the LHA, and NTS, all nuclei with direct
projections to the PVH (Levin, Dunn-Meynell, and Routh, 1999). Further,
dramatic changes in brain glucose levels are linked to changes in food intake
and autonomic motor output (Davis, Wirtshafter, Asin, and Brief, 1981).
Although some of these effects have been shown to be independent of
insulin (Levin, 1991), normally, glucose and insulin levels are intimately
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linked. Thus, in physiological settings an increase of plasma glucose will
also result in an excursion of circulating insulin. Combination tract- tracing
and immunohistochemistry in animals after administration of insulin and
glucose at levels that occur post-prandially identified activated neurons in the
medial parvocellular PVH with projections to the autonomic preganglionic
spinal cord, suggesting their direct involvement with the sympathetic control
of glucose metabolism (Carrasco, Portillo, Larsen, and Vallo, 2001) .
Metabolic hormones: The central nervous system receives information
regarding metabolic status through circulating insulin, leptin, and glucorticoid
levels. Although insulin can affect motor output through its effects on
circulating glucose, several lines of investigation have suggested insulin can
also act directly within the central nervous system to regulate feeding and
metabolism. First, active transport mechanisms have been identified which
carry insulin across the blood-brain barrier (Schwartz, Bergman, Kahn,
Taborsky, Fisher, Sipols, Woods, Steil, and Porte, 1991) and specific insulin
receptors have been located in the brain (Unger, Livingston, and Moss,
1991). Second, these receptors are located within nuclei known to be
involved in energy regulation, particularly the arcuate nucleus (Baskin,
Sipols, Schwartz, and White, 1993; Unger etal., 1991) and
intracerebroventricular injection of insulin reduces food intake and body
weight (Woods, Lotter, McKay, and Porte, 1979). Finally, female mice with a
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specific knock-out disruption of the neuronal insulin receptor gene have
increased feeding and body fat (Bruning, Gautam, Burks, Gillette, Schubert,
Orban, Klein, Krone, Muller-Wieland, and Kahn, 2000).
Central insulin targets neurons in the arcuate nucleus of the hypothalamus
where it appears to affect NPY synthesis and release. For example,
streptozotocin-induced diabetes in rats causes diabetes-induced
hyperphagia and increases hypothalamic NPY expression, both of which are
normalized by insulin treatment (Havel, Hahn, Sindelar, Baskin, Dallman,
Weigle, and Schwartz, 2000; Marks, Waite, and Li, 1993). Further insulin
therapy normalizes food deprivation-induced NPY release into the PVH in a
dose-dependent manner (Sahu, Dube, Phelps, Sninsky, Kalra, and Kalra,
1995). Together these results suggests insulin is a peripheral signal of
metabolic status that is transduced by insulin-sensitive ARH neurons and can
affect food intake.
Insulin has also been shown to modulate the intracellular mechanisms
responsible for transducing leptin signals (Carvalheira, Siloto, Ignacchitti,
Brenelli, Carvalho, Leite, Velloso, Gontijo, and Saad, 2001). Leptin, a protein
product of peripheral adipocytes, is secreted into the circulation to reflect the
level of body fat and the energy balance state of the animal. A deficiency of
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circulating leptin, as seen in both starvation and in ob/ob mice (Campfield,
Smith, Guisez, Devos, and Burn, 1995; Dallman, Akana, Bhatnagar, Bell,
Choi, Chu, Horsley, Levin, Meijer, Soriano, Strack, and Viau, 1999),
stimulates eating at least in part by increasing hypothalamic NPY levels
(Ahima, Kelly, Elmquist, and Flier, 1999). The NPY producing neurons found
in the ARH express leptin receptors (Mercer, Hoggard, Williams, Lawrence,
Hannah, Morgan, and Trayhurn, 1996) and project to the PVH and the LHA
(Broberger, Visser, Kuhar, and Hokfelt, 1999), areas that when injected with
NPY, show eating behaviors (Stanley, and Leibowitz, 1985). Leptin is
expressed with strong diurnal changes reflecting the timing of food intake,
and changes in feeding patterns can change the pattern of leptin expression
(Schoeller, Celia, Sinha, and Caro, 1997). This daily leptin surge may encode
a signal that reflects daily intake and its absence is a potent signal of
starvation (Ahima, 2000). This surge is in addition to the basal secretion of
leptin which encodes for the level of body fat storage (Ahima, 2000).
Whether or not this second pattern of leptin expression is important as a
feedback mechanism leading to a change in drive state is unclear, although
the first is probably important as a starvation signal.
Working in conjunction with insulin and leptin as a peripheral signal of energy
balance are circulating glucocorticoids (GCs). GCs are released from the
adrenal cortex in response to stress or starvation (Dallman et al., 1999) as a
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result of activity in PVH neuroendocrine motor neurons. Data suggest GCs
are important mediators of energy balance, although their specific
mechanisms remain unclear, largely due to their complex inter-relationships
with insulin and leptin. For example, glucocorticoids have been shown to
increase NPY mRNA and food intake in diabetic rats (Strack, Sebastian,
Schwartz, and Dallman, 1995) as well as produce eating in spite of elevated
leptin levels and normal NPY gene expression (Strack et a!., 1995). GCs
also may have a role in the autonomic responses involved in energy balance
as adrenalectomy abolishes the increased insulin response normally seen
after NPY injection (Wisialowski, Parker, Preston, Sainsbury, Kraegen,
Herzog, and Cooney, 2000).
Neuropeptide Y
In the discussion of motor events underlying energy balance we have
described many of the signals of peripheral metabolism status affecting the
central nervous system though activity of NPY in the hypothalamus. In
fact, NPY is a neuropeptide that is integral in homeostatic energy balance.
NPY is a 36 amino acid neurotransmitter that is highly conserved across
diverse species (Larhammar, Blomqvist, and Soderberg, 1993). NPY
belongs to a family of related peptides that includes pancreatic polypetide
and peptide YY, the letter Y derived from the tyrosine residues found at
each end of the hairpin-shaped molecule. NPY is one of the most abundant
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and widely distributed amino acids in the central nervous system. However it
is found in extremely high concentrations in hypothalamic nuclei, particularly
those that have a role in regulating appetite and metabolism. The majority of
NPY in the hypothalamus is produced by those neurons residing in the ARH
which send efferent projections to other hypothalamic nuclei including the
PVH and the LHA, particularly the perifornical part (LHApf, (Broberger et at.,
1998b; Elias etal., 1998b). There are also significant NPY-containing fibers
ascending from the medulla to these same hypothalamic nuclei (Bai,
Yamano, Shiotani, Emson, Smith, Powell, and Tohyama, 1985; Sawchenko,
Swanson, Grzanna, Howe, Bloom, and Polak, 1985). NPY is a powerfully
orexigenic neurotransmitter that has been well characterized in its ability to
increase food intake. When administered directly into the PVH or the LHApf,
NPY elicits voracious feeding while decreasing energy expenditure
(Billington, Briggs, Grace, and Levine, 1991; Morley, 1987; Stanley etal.,
1984; Stanley, Magdalin, Seirafi, Thomas, and Leibowitz, 1993). The fact
that chronic infusions of NPY produces obesity (Stanley, Kyrkouli, Lampert,
and Leibowitz, 1986; Zarjevski, Cusin, Vettor, Rohner-Jeanrenaud, and
Jeanrenaud, 1993) and that gene expression and release of NPY is linked
with an animal’s metabolic state suggests NPY is involved in the regulation of
body weight. In fact, manipulations that increase food intake such as
starvation or diabetes increase NPY synthesis, release, tissue content and
gene expression (Beck et al., 1990; Kalra, Dube, Sahu, Phelps, and Kalra,
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1991; Sahu, Kalra, and Kalra, 1988; Williams, Gill, Lee, Cardoso, Okpere,
and Bloom, 1989). In summary, several mediators of peripheral metabolism
change during starvation; circulating insulin and leptin decrease, while
glucocorticoids increase. These provide afferent information to NPY neurons
which activate the ARH-PVH-LHA neural network which is important for
refeeding after energy deficit.
ANOREXIA
When appetite and food intake are not increased despite the presence of
multiple markers of negative energy balance, it suggests a fault in NPY-
driven mechanisms contributes to the failure to stimulate usual adaptive
feeding responses. The reasons why these mechanisms fail in different
forms of anorexia remain largely unknown. Adaptive feeding responses may
be ineffective because of a faulty feedback system by which NPY synthesis
and release is not activated by falling levels of peripheral metabolic signals.
Conversely, other mechanisms may alter NPY terminal sites thus making
them less responsive to the NPY arising from ARH or hindbrain efferents.
While it is apparent that different types of anorexia occur in a variety of
circumstances and the various mechanisms responsible for different types of
anorexia are no doubt diverse, they all result in a disruption of the normal
feeding response to starvation and so presumably they must converge on a
common set of neural networks that control feeding behavior.
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In order to understand the different mechanisms underlying anorexia it is
useful to begin by categorizing anorexia types based on available
experimental evidence of hypothalamic NPY profiles. Because NPY has
become one of the best characterized feeding systems, it can also serve as
an effective tool by which to probe neural mechanisms underlying different
types of anorexia. In fact, by using hypothalamic NPY profiles anorexia can
by divided, at least at a first approximation, into two types.
Anorexia characterized by continued sensitivity to NPY
Some anorexia types appear to result from a pathological augmentation of a
negative feedback system that normally decreases NPY activity in the ARH.
Decreased NPY may occur because of abnormally increased leptin levels or
from mechanisms in NPY neurons that utilize leptin-like signals. These
types of anorexia are often associated with inflammation of infectious agents
that stimulate macrophages and monocytes to produce cytokines. Cytokines
orchestrate the host’s inflammation response through the production of
acute-phase proteins and the mobilization of lymphocytes, while increasing
blood flow to improve vascular permability (Gabay, and Kushner, 1999).
Elevated cytokine levels elicit neuroendocrine and metabolic responses
leading to fever, somnolence and anorexia (Gabay etal., 1999). In fact, the
levels of circulating cytokines are tightly correlated with anorexia and its
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accompanying weight loss in many circumstances (Aleman, Santolaria,
Batista, de La Vega, Gonzalez-Reimers, Milena, Llanos, and Gomez-Sirvent,
2002; Ballinger, Kelly, Hallyburton, Besser, and Farthing, 1998; Pereira, and
Dinarello, 1994; Plata-Salaman, 1996; Takabatake, Nakamura, Abe, Hino,
Saito, Yuki, Kato, and Tomoike, 1999; Verbon, Juffermans, Van Deventer,
Speelman, Van Deutekom, and Van Der Poll, 1999).
The regulation of brain cytokines during peripheral inflammation or infection
is unclear. Cytokines themselves are produced and released by microglia
and the endothelial cells of the circumventricular organs or vasculature of the
brain, or alternatively cytokines may be transported across the blood brain
barrier (Maness, Kastin, and Banks, 1998) or by neural signaling provided by
either vagal or sympathetic afferents (Goldbach, Roth, and Zeisberger,
1997). Two pro-inflammatory cytokines, tumor necrosis factor-a (TNF-a) and
interleukin-1 (IL), induce both their own expression and that of other
cytokines including IL-6, IL-11, ciliary neurotrophin factor (CNTF), and
interferon-y. Both TNF-a and IL-1 have been proposed as mediators in the
anorexic response to disease. Chronic administration of TNF-a or IL-1
increases leptin gene expression, stimulates peripheral leptin release
(Kirchgessner, Uysal, Wiesbrock, Marino, and Hotamisligil, 1997) and
produces a dose-dependant increase in serum leptin levels in both
experimental animals and in humans (Grunfeld, Pang, Shigenaga, Jensen,
Lallone, Friedman, and Feingold, 1996; Sarraf, Frederich, Turner, Ma,
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Jaskowiak, Rivet, Flier, Lowell, Fraker, and Alexander, 1997; Zumbach,
Boehme, Wahl, Stremmel, Ziegler, and Nawroth, 1997). In this manner, one
potential mechanism of how cytokines affect food intake is the pathological
augmentation of the negative feedback system normally engaged by leptin,
which reduces feeding, body weight, and masks the compensatory
mechanisms that normally are initiated by these changes. However, clinical
conditions where an inflammatory response is present with both weight loss
and increased circulating leptin are surprisingly rare. In inflammatory bowel
conditions with elevated proinflammatory cytokines (Ballinger etal., 1998;
MacDermott, 1996) patients do tend to have higher circulating leptin than
matched controls (Bannerman, Davidson, Conway, Culley, Aldhous, and
Ghosh, 2001). Likewise, patients suffering from chronic renal failure, where
chronic inflammation is a common feature (Bergstrom, 1999; Merabet,
Dagogo-Jack, Coyne, Klein, Santiago, Hmiel, and Landt, 1997) also appear
to have elevated blood leptin levels. However, weight-losing patients with
chronic obstructive pulmonary disorder have increased circulation levels of
TNF while serum leptin levels in these patients are not elevated over those in
healthy controls (de Godoy, Donahoe, Calhoun, Mancino, and Rogers, 1996;
Di Francia, Barbier, Mege, and Orehek, 1994; Takabatake etal., 1999).
Furthermore, in patients with advanced lung cancer or advanced acquired
immunodeficiency syndrome (AIDS), leptin levels were inversely related to
the intensity of the inflammatory response (Aleman et al., 2002; Ballinger et
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a/., 1998). Even severe inflammation resulting from sepsis did not elevate
leptin levels in humans (Carlson, Saeed, Little, and Irving, 1999). When
considered together, it appears that even intense acute inflammatory
responses that produce high levels of cytokines do not necessarily result in
high serum leptin levels in many cases of anorexia accompanying disease.
Therefore it is unlikely that the anorexia seen in these cases is due to a
dysregulation in leptin production or release.
Alternatively, in conditions of inflammation it is possible that the elevated
cytokines engage the same catabolic effector pathways normally used by
leptin in the hypothalamus, because leptin and cytokines are structurally
similar. Furthermore, the leptin receptor is a member of the class 1 cytokine
family known to signal through the Janus kinase/signal transducer and
activator of transcription (STAT) pathway (Tartaglia, Dembski, Weng, Deng,
Culpepper, Devos, Richards, Campfield, Clark, Deeds, and et al., 1995).
The STATs, once activated, dimerize and translocate to the nucleus where
they affect gene expression (Darnell, 1997). A peripheral injection of either
leptin or TNF-a stimulates STAT3 tyrosine phosphorylation in the
hypothalamus, and injection of both leads to a synergistic increase in
phosphorylation (Rizk, Stammsen, Preibisch, and Eckel, 2001), suggesting
the strong possibility of common signaling pathways. In fact, the glycoprotein
130 signal-transducing subunit of the IL-6 type receptors is a homolog of the
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leptin receptor (Baumann, Morelia, White, Dembski, Bailon, Kim, Lai, and
Tartaglia, 1996). Evidence supports this hypothesis in experimental animals.
For example, icv injection of IL-1 induces anorexia and may elicit some of the
same central effects on feeding pathways as those seen after leptin injection
(Gayle, Ilyin, and Plata-Salaman, 1997). CNTF also potently elicits
decreased food intake and body weight when injected either systemically or
into the brain, and can engage at least some of the same neural signaling
pathways that are activated by leptin (Pu, Dhillon, Moldawer, Kalra, and
Kalra, 2000; Xu, Dube, Kalra, Farmerie, Kaibara, Moldawer, Martin, and
Kalra, 1998). In particular, CNTF anorexia develops despite reduced leptin
levels (Lambert, Anderson, Sleeman, Wong, Tan, Hijarunguru, Corcoran,
Murray, Thabet, Yancopoulos, and Wiegand, 2001; Pu etal., 2000). Critical
to this idea however, is that in experimental animals all cytokine-related types
of anorexia demonstrate reduced hypothalamic NPY gene expression and
release, and they all respond to exogenous NPY administration with
increased food intake (Inui, 1999; Pu etal., 2000; Turrin, Flynn, and Plata-
Salaman, 1999).
Anorexia characterized by reduced sensitivity to NPY
Neural mechanisms that render the hypothalamus insensitive to NPY
characterize the second category of anorexia. This includes the anorexia
that develops with certain types of tumors, colitis, aging and cholestasis.
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(Ballinger, Williams, Corder, El-Haj, and Farthing, 2001; Blanton, Horwitz,
Blevins, Hamilton, Hernandez, and McDonald, 2001; Chance,
Balasubramaniam, Thompson, Mohapatra, Ramo, and Fischer, 1996;
Jensen, Blume, Mikkelsen, Larsen, Jensen, Holst, and Madsen, 1998;
Makino, Asaba, Nishiyama, and Hashimoto, 1999; Rioux, Le, and Swain,
2001). Anorexic animals with tumors have lowered leptin levels and
increased NPY gene expression in the ARH (Chance etal., 1996; Chance,
Sheriff, Kasckow, Regmi, and Balasubramaniam, 1998; Jensen etal., 1998).
Identical profiles are found in anorexic rats after experimentally-induced
colitis (Ballinger etal., 2001), in anorexia that accompanies dehydration
(O'Shea, and Gundlach, 1995; Watts et at., 1999) and in anorexia that
follows psychological stress or normal aging (Blanton eta!., 2001; Makino et
al., 1999; Makino, Baker, Smith, and Gold, 2000). Critically, unlike our first
anorexia category characterized by continued sensitive to NPY, anorexia in
these circumstances remains resistant to the stimulatory feeding effects of
hypothalamic NPY administration (Ballinger etal., 2001; Blanton etal., 2001;
Chance etal., 1996; Inui, 1999; Rioux et al., 2001). Additionally,
underweight patients with AIDS, obstructive pulmonary disease,
inflammatory bowel disease, and anorexia nervosa have lower circulating
leptin levels (Ballinger et al., 1998; Grunfeld etal., 1996; Kaye, Klump, Frank,
and Strober, 2000; Polito, Fabbri, Ferro-Luzzi, Cuzzolaro, Censi, Ciarapica,
Fabbrini, and Giannini, 2000; Takabatake etal., 1999), suggesting elevated
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NPY gene expression, if human ARH NPY neurons respond to leptin as in
experimental animals. Although patients with AIDS and anorexia nervosa
have elevated cerebral spinal fluid (CSF) NPY levels, direct post-mortem
analysis of the human ARH has yet to be directly examined in these
illnesses. However, of interest are reports of post-mortem analysis of
humans with illness-associated anorexia finding elevated NPY gene
expression in the hypothalamus (Corder, Pralong, Muller, and Gaillard, 1990;
Goldstone, Unmehopa, Bloom, and Swaab, 2002).
Collectively these data suggest that the reduced drive to eat in these
circumstances of anorexia is not due to an inhibition of peripheral metabolic
signal-sensitive NPY mechanisms that usually stimulate appetite.
Alternatively, it appears there is an increase level of activity in these circuits
and rather the output of these mechanisms is inhibited. The inhibitory
mechanisms responsible are not known, but some forms of anorexia can be
partly reversed by manipulating melanocortin or corticotropin releasing
hormone receptors (Lawrence, and Rothwell, 2001; Smagin, Howell,
Redmann, Ryan, and Harris, 1999; Wisse, Frayo, Schwartz, and Cummings,
2001) implicating a potential role for the ligands of these receptors. Still, the
central localization of such inhibitory substrates or their interaction with
stimulatory circuits is not clear.
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We have addressed these questions with a series of experiments designed
to explore the organization of central networks underlying the anorexia that
develops with dehydration (DE). DE is a classic homeostatic challenge that
results in a series of adaptive responses aimed to normalize imbalances in
volume or osmolality. Reciprocally linked osmosensitive sites in the lamina
terminalis, including the subfornical organ (SFO), the median preoptic
nucleus (MnPO) and the organum vasculosum of the lamina terminalis
(OVLT), rigidly monitor the fluid compartment and respond to perturbations
through projections to a network of forebrain cell groups which contribute to
the homeostatic response, of which the PVH, LHA and fusiform part of the
bed nucleus of the stria terminalis (BNSTfus) appear to play prominent roles
(Kelly, and Watts, 1996; Swanson, 1991; Swanson, and Sawchenko, 1983;
Watts, Kelly, and Sanchez-Watts, 1995a; Watts, and Sanchez-Watts, 1995b;
Watts et al., 1999). In the face of a disruption of body fluid homeostasis, a
triad of neuroendocrine, autonomic and behavioral motor events are
employed to protect and normalize water balance. First, elevated plasma
vasopressin (AVP) secretion serves to increase water retention from the
kidneys. Second, hemodynamic changes are activated to optimize the
distribution of water to tissues and maintain blood pressure and third,
spontaneous circadian driven food intake is reduced to limit the addition of
osmoles into the already compromised fluid compartment (Dicker, and Nunn,
1957; Johnson, and Gross, 1993). By limiting the amount of food in the gut,
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anorexia also helps to preserve fluid balance by slowing digestion and
allowing fluid to be redirected towards the more vital cellular fluid
compartment.
For our experiments, animals were dehydrated by providing a hypertonic
2.5% saline solution in place of their regular drinking water for up to five
days. As animals become progressively dehydrated by drinking hypertonic
saline, they exhibit a profound anorexia that can be reversed quickly by
restoring water access. Thus, DE-anorexia provides a useful model to probe
underlying circuits and mechanisms that might be responsible for inhibiting
stimulatory feeding behaviors, particularly when compared to other models of
anorexia. For example, some models induce anorexia by implanting tumors,
injecting the bowel with acids, or providing the animal with psychogenic
stress. These approaches generate multiple input stimuli, some of which
may be anorexigenic while others may not, making it difficult to discriminate
between cause and effect. In addition, it is often difficult or impossible to
reverse various models of anorexia. In contrast, DE-anorexia has several
utilitarian properties that effectively constrain its effector mechanisms. The
anorexia develops within two days, simply by providing animals with
hypertonic saline to drink. Drinking hypertonic saline results in predominately
intracellular DE, thus effectively limiting potential signaling mechanisms to
that of a well-characterized physiological stimulus (Watts, 1992; Watts, and
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Sanchez-Watts, 1995c; Watts etal., 1999). Finally, DE-anorexia can be
reversed within minutes simply by returning drinking water to the animals
(Watts, 1999). Together these characteristics permit effective investigation
of the neural networks that might stimulate, inhibit, or switch ingestive
behaviors.
The hormonal and neural profile of DE-anorexia places it convincingly into
the type of anorexia we have characterized as being resistant to the feeding
stimulatory effects of NPY. Dehydrated animals demonstrate body weight
loss, diminished circulating leptin and insulin levels, and increased circulating
blood glucocorticoids, in addition to an increased NPY gene expression the
ARH (Watts et al., 1999), all signals normally associated with increased food
intake. In fact, euydrated (EU) control animals food restricted to match the
food eaten by DE animals show an identical hormonal profile and
demonstrate an appreciable appetite. However, DE animals remain anorexic
until given back water to drink (Salter and Watts, 2003; Watts et al., 1999).
The focus of this thesis is to test the overall hypothesis that dehydration
results in a post-synaptic inhibition of NPY-associated feeding mechanisms.
A series of experiments are presented using DE-anorexia as a model for
identifying the underlying circuits and mechanisms that might be responsible
for inhibiting NPY effects on feeding. We begin by probing the specificity of
the inhibition that accompanies DE by examining the viability of different
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motor responses to metabolic challenge. Next, we explore neural activation
patterns for areas that might be inhibited by DE. We then test the integrity of
the feeding response in DE animals through central injections of orexigenic
agents. As such, we begin to address fundamental questions that remain in
ingestive behaviors, that is how appetite can be initiated and turned off, and
thus inform study of both obesity and anorexia. In a broader sense, the
experiments in this thesis are designed to address the pathophysiology of
constituent neural circuits and transmitters in the disequilibrium between
energy intake and expenditure in both obesity and anorexia. Therefore, work
done here to clarify the neural circuits and their mechanistic attributes is
critical for understanding behaviors regulating energy balance, specifically
those that control appetite and how the disruption of these neural systems
contributes to anorexia and body wasting.
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CHAPTER TWO
DEHYDRATION ANOREXIA AND GLUCOREGULATION
INTRODUCTION
DE initiates a series of adaptive responses that target gastrointestinal
function in an effort to conserve water and limit the addition of osmoles to an
already compromised fluid compartment. Thus, salivation, gut motility, and
food intake are all reduced in an attempt to resolve or minimize fluid
perturbations at the expense of energy balance (Dicker etal., 1957;
Flanagan, Dohanics, Verbalis, and Strieker, 1992; Watts etal., 1999).
Therefore, DE-anorexia offers a useful paradigm for investigating how the
normal compensatory mechanisms used to trigger feeding during negative
energy balance can be inhibited.
We have already demonstrated that both dehydrated animals and animals
that are food restricted (FR) to match the intake of anorexic rats show the
same attributes of negative energy balance (Watts et al., 1999). These
include body weight loss, diminished circulating leptin and insulin, and
increased blood glucocorticoid concentrations and neuropeptide Y (NPY)
gene expression in the arcuate nucleus of the hypothalamus (ARH).
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Normally these neural and endocrine processes stimulate compensatory
feeding mechanisms aimed at increasing caloric intake to match expenditure.
In this way, weight loss triggers hunger to restore body energy stores through
food intake. Elevated glucocorticoids and decreased insulin levels can
independently stimulate food intake (Baskin, Figlewicz Lattemann, Seeley,
Woods, Porte, and Schwartz, 1999; Solano, and Jacobson, 1999) while
falling plasma leptin levels after starvation (Dallman et al., 1999; Friedman,
and Halaas, 1998) or the deficiency of leptin in ob/ob mice (Campfield,
Smith, and Burn, 1996) both stimulate eating.
Feeding behavior is mediated in certain circumstances by the increased
activity of those NPY-producing neurons in the ARFI (Ahima et al., 1999;
Sahu et al., 1988) that express leptin receptors and project to the
paraventricular nucleus of the hypothalamus (PVFI) and the lateral
hypothalamic area (LFIA) (Broberger, 1999; Elias et al., 1998a). NPY
injections in the PVH or LHA elicit robust feeding (Leibowitz et al., 1988;
Stanley etal., 1984; Stanley et al., 1993) while chronic infusions of NPY
leads to hyperphagia and obesity (Zarjevski etal., 1993). Based on their
neuropeptide and endocrine profile, we have proposed that in DE-anorexic
animals, some component of these NPY-mediated compensatory
mechanisms is inhibited until released by subsequent water intake (Watts,
2000).
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The present study was designed to determine whether DE-anorexic rats
show reduced feeding responses to two challenges that invoke feeding
responses thought to involve NPY-mediated mechanisms: 2-deoxy-D-
glucose (2DG) and overnight starvation. NPY appears to be a key mediator
of fasting-induced hyperphagia in that food deprivation increases NPY mRNA
in the ARH (Dallman et al., 1999), and elevates both NPY levels and release
in the PVH (Kalra etal., 1991; Sahu etal., 1988; Yoshihara, Honma, and
Honma, 1996). 2DG is a glucose analogue that leads to cytoglucopenia by
competitively inhibiting glucose utilization. Evidence implicates NPY in the
feeding response that follows either central or peripheral administration of
2DG (Akabayashi, Zaia, Silva, Chae, and Leibowitz, 1993; Giraudo, Grace,
Billington, and Levine, 1999; He, White, Edwards, and Martin, 1998; Minami,
Kamegai, Sugihara, Suzuki, Higuchi, and Wakabayashi, 1995). 2DG feeding
is mediated by central catecholamine neurons (Li, and Ritter, 2004; Ritter,
Dinh, Sanders, and Pedrow, 2001b; Ritter, Dinh, and Zhang, 2000; Ritter et
al., 1998) which colocalize NPY and project to the PVH (Sawchenko etal.,
1985). Immunoneutralization of NPY in the PVH impairs 2DG feeding (He et
al., 1998) while 2DG-induced glucoprivation increases Fos expression in
ARH and hindbrain neurons that express NPY (Li et al., 2004; Minami et al.,
1995).
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2DG offers a further advantage as an experimental tool for delineating the
neural circuits underlying anorexia because, in addition to stimulating
feeding, it rapidly elicits two other motor responses: sympathetic activation of
epinephrine secretion from the adrenal medulla, which leads to
hyperglycemia (Scheurink, and Ritter, 1993); and corticosterone secretion
from the adrenal cortex (Weidenfeld, Corcos, Wohlman, and Feldman, 1994),
which is driven by increased activation of neuroendocrine CRH neurons in
the PVH. Together, these behavioral, autonomic, and neuroendocrine motor
events serve to replenish and redistribute metabolic fuels. Examining how
other non-behavioral motor control systems function during DE should
provide a broader view of how adaptive neural mechanisms function in
anorexia.
MATERIALS AND METHODS
ANIMALS AND PROCEDURES
Adult male Sprague-Dawley rats weighing 235-260 grams were obtained
from Harlan Laboratories and housed in suspended Plexiglas cages with
sanitized wood chips. They were maintained in a temperature controlled
room on a 12h:12h light:dark schedule with lights on at 06.00h. Rats were
provided with continuous access to food (Teklad rodent diet 8604) and water
throughout experiment, except where stated. In some animals, drinking water
33
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was replaced with 2.5% saline solution for up to 5 days; in others, the
amount of food available was restricted to match that eaten by DE animals
(54). We have previously shown that five days of drinking hypertonic saline
increases plasma osmolality by approximately 6% (Watts et al., 1995a; Watts
etal., 1999). Body weights and nocturnal food intake were measured daily
throughout the experiment. All procedures have been approved by the local
IACUC.
SURGICAL PROCEDURES
Rats were handled daily for approximately 4 days before any surgical
intervention and daily thereafter. On the fourth day after arrival, rats
designated for 2DG injections were anesthetized with an intramuscular
100pl/kg injection of a 50% solution of Ketamine (100mg/ml) plus xylazine
(20mg/ml) and sterile intra-atrial catheters inserted by way of the external
jugular vein. Catheters were threaded subcutaneously to the dorsal surface,
exteriorized between the scapulae, and then sutured in place. Catheters
were flushed daily with sterile heparinized 0.9% saline. Animals were allowed
to recover to pre-surgical weight before further manipulation. Only rats with
stable weights gains and consistent nocturnal intakes were included in the
study.
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OVERNIGHT STARVATION
Five groups of animals were used in this experiment. Food intakes were
measured in all groups for three days prior to testing. Drinking water was
replaced in two groups with 2.5% saline for either 3 (DE-3d) or 5 days (DE-
5d). Body weights and food intake were then measured twice daily between
08.00-09.O O h and 16.00-17.OO h. Food was removed from all cages at 17.00h
on the evening of the final night of saline. The following morning at 08.O O h a
measured amount of food was returned and the food remaining in the cage
measured to the nearest 0.1 gram each hour for a total of four hours. At the
conclusion of the feeding test, saline was replaced with drinking water and
food intake measured after a further hour.
A food restriction (FR) schedule was provided for two other groups of
animals maintained on drinking water. Animals were weight-matched to
animals DE for 3 (FR-3d) or 5 days (FR-5d). They were then given an
amount of food at the beginning of each light and dark period equal to that
eaten by DE animals (Watts et al., 1999). The amount of food was
calculated for each rat as a percentage of the food eaten per 100g of mean
body weight (for two days before beginning the food restriction). On the
evening of the second or the fourth day of FR, food was removed completely.
Food was returned to animals the following morning and intake measured
each hour for the next four hours. A fifth group of euhydrated (EU) animals
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
was allowed continuous access to food (ad-lib) and water. Food intake was
measured in this group for four hours the morning following an overnight fast.
RESPONSES TO 2-DEOXY-D-GLUCOSE
Four experiments were performed to determine the effect of DE on the
responses to 2DG. Experiment 1 established a dose-feeding response curve
for 2DG. Experiment 2 determined the effect of 2 doses of 2DG given 7 days
apart to test the validity of using each animal as its own control before and
during DE. Experiment 3 then determined the effects of DE on feeding
responses to 2DG. In Experiment 4, we measured the effects of DE on the
plasma glucose and corticosterone responses to 2DG.
All feeding tests were conducted between 08.O O h and 13.O O h as follows. At
the beginning of the experiment all food and sawdust was removed from
cages, animals were weighed and a measured amount of food placed in the
test cage for approximately one hour. Equal volumetric doses of vehicle or
2DG (0.1 ml/kg) were then injected into the jugular catheter. Some rats in
Experiments 3 and 4 were given a subcutaneous 2DG injection because of
blocked jugular catheters. There was no significant difference in response
between catheter and subcutaneous injections, so these data were pooled.
36
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
Following the injection, food consumption was measured by weighing the
food remaining in each cage to the nearest 0.1 gram each hour for the next
four hours.
Experiment 1
To establish appropriate doses of 2DG for investigating the effects of DE on
the feeding response to 2DG, animals maintained with continuous access to
water were injected with either vehicle (0.9% saline) or one of four doses of
2DG; 50 mg/kg, 100 mg/kg, 200 mg/kg, and 250 mg/kg. Food intake was
then measured as just described.
Experiment 2
One group of animals was tested three times over the next 9 days for food
intake after injections of either vehicle or 2DG. On day one, vehicle injections
were given to measure baseline food consumption in the test cage. The
following day each animal was given a 200mg/kg 2DG injection. Food intake
was again measured on day 9 after a 200mg/kg 2DG injection.
Experiment 3
Rats were divided into three groups and tested three times over the next 9
days for food intake after injections of either vehicle or 2DG. Vehicle
injections were given on day one to all animals to establish baseline food
37
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
consumption in the test cage. The following day (day 2) each animal was
given a vehicle, 200, or 250 mg/kg 2DG injection. On day four, drinking water
was replaced with 2.5% saline. Food intake was again measured on day 9,
the fifth day of DE after a vehicle, 200, or 250 mg/kg 2DG injection. Each
animal received the same treatment on days 2 and 9. At the conclusion of
the four-hour feeding test, drinking water was returned to all DE animals and
food intake measured during the following hour.
Experiment 4
The design of this experiment was virtually identical to Experiment 3 except
that blood samples were taken from the jugular catheter for plasma glucose
and corticosterone determinations. Animals were tested three times after
equal volumetric injections of 0.9% saline vehicle or 2DG (0.1 ml/kg). Venous
blood samples (150pl) were collected from the jugular catheters of all
animals immediately before and 15, 30, 60, and 120 minutes after injection.
Food was removed for 2 hours before baseline blood collection and not
returned until after the final blood sample.
On day one, all animals received an injection of vehicle to determine baseline
responses; on day three, animals received an intra-atrial injection of vehicle
or 200 mg/kg 2DG. Drinking water was replaced with 2.5% saline at 12.00h
on the following day. On the morning of the fifth day of DE, rats were injected
38
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
with vehicle or 200 mg/kg of 2DG. Drinking water and food was replaced at
the conclusion of the blood sampling. Only animals that exhibited feeding
response of more than 3g to drinking water were included in the study.
Blood samples were immediately placed on iced, centrifuged, plasma
removed, and stored at20°C until assayed. Plasma glucose concentration
was assayed using a YSI 2700 auto-analyzer (Yellow Springs, OH). Plasma
corticosterone concentrations were determined by double antibody
radioimmunoassay (Tanimura, Sanchez-Watts, and Watts, 1998) using a
commercial available kit (ICN Pharmaceuticals). All samples were assayed in
duplicate in single assays. Internal controls were within appropriate ranges.
Food intake meaured • 1-4h
i ,
i 1-4h
! I
« 1-4h
i
i I
1h
| 4-5 days j 1 day | 2 days j 5 days | 4 hours |
Cannulate vehicle
(2DG group)
2-DG
(2DG group)
or vehicle
(control group)
Dehydration 2-DG
(2DG group)
or vehicle
(control group)
Water returned
Figure 2.1: Timeline for experiments on EU and DE animals given 2DG.
Animals were given the first injection of VEH or 2DG 4-5 days following
surgery. Each animal then received an injection of 2DG or VEH two days
later in a counterbalanced design. Animals were placed into groups, two
groups being DE and one group continuing on ad lib water intake. At the
end of the DE period (5 days) all DE animals were tested for their feeding
response to either a second injection of 2DG or VEH, and EU animals were
tested for their response to a second injection of 2DG. DE animals were
given water back at the conclusion of the four hour test.
39
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
STATISTICS
Data are expressed as means ± SEM. Changes in body weight during DE,
food intake suppression, food intake after overnight starvation, and amount
eaten after return of water were compared between groups using one-way
ANOVA. For analysis of plasma glucose and corticosterone responses,
incremental increases were calculated for each animal by subtracting the
maximum concentration attained from the pre-injection value. Baseline EU
tests were compared to EU 2DG tests in experimental animals using ANOVA
with repeated measures at each time point. Data from EU and DE animals
were compared using a 2 x 2 repeated-measure ANOVA. Bonferrroni's
multiple comparison test was used to measure individual differences. The
critical level for significance was set at p<0.05 for all comparisons.
RESULTS
FEEDING RESPONSES TO OVERNIGHT STARVATION
Overnight starvation elicited significantly different amounts of compensatory
feeding in DE, FR, and ad lib groups (at hour 4, F {df 4,25} =45.92, P<
0.001). Figure 2.2A shows that the amount of food eaten after 4 hours by FR-
5d animals was significantly greater than that consumed by ad lib animals
(p<0.05). However, both groups of DE animals ate significantly less after
40
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
overnight starvation than both FR and al lib animals, with the DE-5 animals
eating significantly less than the DE-3 animals (p< 0.01). This represented a
60% and 80% reduction compared to that displayed by the ad lib fed EU
group (Fig. 2.2B). However, all DE animals displayed a robust eating
response during the hour after access to drinking water was restored (Fig.
2.2A), the size of which was not significantly different between the two DE
groups.
FEEDING RESPONSES TO 2-DEOXY-D-GLUCOSE
Changes in body weight over the course of the feeding tests are shown in
Table 2.1. There was no significant difference in rate of weight gain among
the animals in the three treatment groups before or during the testing period.
At the end of DE, rats in each treatment group lost a similar amount of body
weight and exhibited equivalent anorexia (Table 2.1).
Experiment 1
Figure 2.3 shows the amount of food eaten after injection of increasing doses
of 2DG. 50 mg/kg or 100 mg/kg 2DG elicited a feeding response that was no
different than vehicle at all time points except at hour 4, when the response
to 100 mg/kg was significantly greater (p<0.05) than vehicle or 50 mg/kg
(Fig. 2.3B). After either 200 or 250 mg/kg 2DG animals ate significantly more
41
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
water returned to
dehydrated animals
Figure 2.2: Hypertonic saline ingestion (DE) inhibits feeding in response to
overnight starvation.
Mean (± SEM) cumulative weight of food eaten after overnight starvation A)
following 5 days of DE (open triangles), 3-day DE (open squares), ad lib
(open circles), 3-day food restriction (FR; closed squares), and 5-day FR
(closed triangles). When water was returned to DE animals there was a
robust eating response measured 1 hour later (dashed lines). B) Mean (±
SEM) total weight of food eaten 4 hours after food replacement by FR (solid
line) or DE (dashed line) animals after overnight starvation and various times
of FR or DE. Data are expressed as a percentage of the response shown by
ad-lib fed animals (100%). See text for levels of statistical significance.
42
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
2DG
vehicle 200 mg/kg 250 mg/kg
Body weight
Beginning, g 248.5
±
4.6 237.1
±
2.2 260.5
+
6.9
Gain, g/day 3.2
+
0.4 2.6
+
0.4 3.5
+
0.4
Decrease with DE,% 13.7
+
2.0 14.8
+
1.3 16.6
+
1.6
Nocturnal food intake
Baseline (2 day avg),g 17.8 ± 0.6 18.6 ± 0.6 18.2 ± 0.4
Decrease with DE,% 59.7 ± 4.6 65.8 ± 4.8 64.9 ± 4.2
Table 2.1: The effects of drinking 2.5% saline on body weights and mean
nocturnal food intake.
There were no significant differences between each treatment group in either
the mean (± S.E.M.) body weights at the start of the experiment, the rate of
increase before DE, or the decline in body weight after DE. Similarly, DE was
associated with a similar decrease in mean (± S.E.M.) nocturnal food intake
in all three groups of animals.
than vehicle at all times (p<0.01). Feeding responses to vehicle injection and
50 mg/kg of 2 DG were indistinguishable. Based on the results of this
experiment, 200 or 250mg/kg were used to test the effects of DE on feeding
responses in subsequent experiments.
Experiment 2
EU animals given two injections of 200mg/kg 2DG separated by 7 days
responded to the second injection in a manner indistinguishable from the first
(Fig. 2.4). One-way repeated measure ANOVA indicated a significant effect
43
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
A B
5.0
4.0
3.0
Q )
ro
T3
§ 2.0
1.0
0.0
T
2 4
T
Time (h)
0 50 100 200 250
Dose 2 DG (mg/kg)
Figure 2.3: The feeding response to various doses of 2-deoxyglucose
injections.
A), Mean cumulative intake (± SEM) each hour for four hours after injection
of vehicle (open circles), 50 (closed squares), 100 (closed triangles), 200
(closed circles) or 250 (closed diamonds) mg/kg of 2-deoxyglucose (2DG).
B), Relationship between dose of 2 DG and food eaten 2 hour after injection.
See text for levels of statistical significance.
44
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
A B
s
©
j*:
ro
TJ
o
o
4.0
3.5
3.0
2.0
0.5
0.0
0 1 2 3 4
r T
0 1 2 3 4
Time (h)
0 1 2 3 4
Figure 2.4: The feeding response to doses of 2-deoxyglucose injections
repeated at 7 day intervals.
A) Mean cumulative intake (± SEM) each hour for four hours after injection of
vehicle (open circles), or (B and C) 200 mg/kg 2DG. The second dose of
2DG in C) was given 7 days after the first B).
of 2DG on food intake (F {df 2,12} = 16.33, P< 0.001). Both the first and
second dose of 2DG elicited significantly more food intake than vehicle at all
time points (p<0.01). However, the amount of food intake eaten after the first
45
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
injection was not significantly different from the amount of food intake after
the second injection at any time.
Experiment 3
Injection of vehicle did not elicit a significant eating response in any of the
three groups of animals (Fig 2.5A), but after administration of either 200
mg/kg or 250 mg/kg of 2DG these animals ate significantly more (Fig. 2.5B; F
{df 2,37} = 48.33, p<0.001). Figure 2.5C shows DE significantly reduced the
ability of 2DG to elicit food intake. Two-way repeated measure ANOVA
revealed a main effect of DE (F {df 2, 25}= 44.25, P<0.001) and an
interaction effect between 2DG and DE (F {df 2, 25} = 12.3, P<0.001).
Finally, all DE animals ate similar amounts of food when drinking water was
returned at the end of the 2DG-feeding test, regardless of whether they were
injected with vehicle or one of the two doses of 2DG four hours previously
(Fig. 2.5D).
Experiment 4
Effects of DE on the Responses of Plasma Glucose Concentrations to 2DG.
Plasma glucose results are presented in Table 2.2 and Figure 2.6. Vehicle
injections did not significantly increase plasma glucose concentrations in any
treatment group on day 1 (Fig. 2.6A). On day 3, animals were injected with
46
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
either vehicle or 200mg/kg 2DG (Fig. 2.6B). There was no significant
response to vehicle, but 2DG elicited a significant increase in plasma glucose
from pre-injection values (F {df 4,35} = 13.25, p < 0.001). On day 9, pre-
A B D
0 )
( 0
T 3
O
O
Li_
5.0
4.5
4.0
3.5
3.0
2.5
2.0
0.5
r
0.0
0 1 0 1 2 3 4
Time (h)
0 12 3 4
5.0
4.5
4.0
^ 3.5
3
i? 3.0
C O
~ a 2.5
o
o
^ 2.0
1.5
1.0
0.5
0.0
Water Back
(1 hour)
Figure 2.5: The effects of 5 days of hypertonic saline ingestion on the
feeding response to 2-deoxyglucose injections.
Mean (± SEM) cumulative food intake in the same group of animals after
injection of vehicle (open symbols), 200 mg/kg 2DG (squares) or 250 mg/kg
2DG (triangles). Food intake is shown after injection of vehicle (A), 2DG
given 1 day later (B), and 2DG given after 5 days of hypertonic saline
ingestion (C). Food intake is also shown in animals given vehicle injection
using the same time schedule (B and C, open circles). (D) illustrates the
mean (± SEM) food intake of the same animals shown in C) 1 hour after
water was returned at the conclusion of the 4 hour 2DG test.
47
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injection plasma glucose concentrations were no different to pre-injection
values on days 1 and 3 (Table 2.2). After injection of vehicle or 2DG, mean
plasma glucose concentrations in DE animals were again unaffected by
vehicle injection, but were significantly increased by 2DG from pre-injection
values (Fig. 2.6C; F {df 4,33} = 9.90, p < 0.001). Two-way ANOVA with
repeated measures indicated that 2DG administration was a significant main
effect at all time points after injection (at 60 minutes, F {df 1,10} = 85.78,
p<0.001). However, DE was not significant as either a main or an interactive
effect at any time point measured. DE did not significantly alter the
incremental increase in glucose elevation elicited by 2DG from that seen in
EU animals at any time point. Additionally, there was no significant difference
in changes of plasma glucose after vehicle injection between animals in the
EU and DE states.
Although DE did not significantly alter the magnitude of the plasma glucose
response to 2DG, there was an apparent difference in the kinetics of the
response. Peak mean glucose concentrations were seen at 15 minutes on
day 3 (EU), but at 120 minutes on day 9 (DE). This was partly due to very
high values in the 15min samples from two animals on day 3. Removing
these values reduced the variance of the mean at 15mins in such a way that
peak values were now seen at 60mins in these animals.
48
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Glucose (mg/dl) Corticosterone (ng/ml)
Day 1 (euhydration)
Vehicle
Pre 104.6 ± 3.0 67.3
+
19.3
Max 112.0 ± 3.0 91.2 ±
31.1
Vehicle
Day 3 (euhydration)
Pre 112.0 ± 4.3 95.9
+
27.8
Max 114.3 ± 5.6 (38.3
+ 18.4)
2-DG
Pre 105.5 ± 1.2 72.0 ± 26.0
Max 270.0 ± 34.3 455.8 ± 54.0
Vehicle
Day 9 (dehydration)
Pre 98.5 ± 11.0 182.0
+
22.6
Max 113.5 ± 25.9 213.6
±
36.6
2-DG
Pre 106.4 ± 4.6 223.0 ± 62.0
Max 246.3 ± 29.6 480.4 ± 41.4
Table 2.2: The effects of drinking 2.5% saline on plasma glucose and
corticosterone responses to injections of vehicle or 2-deoxy-D-glucose.
Injection of vehicle (VEH; 0.9% saline) did not increase mean (± S.E.M.)
plasma glucose and corticosterone either before (Days 1 and 3 EU) or after
drinking 2.5% saline (DE; Day 9). Similarly, 2-deoxy-D-glucose (2DG;
200mg/kg) injections significantly increased plasma glucose and
corticosterone concentrations in all treatment groups. Pre-injection values
(pre) were determined on blood samples taken before injection of VEH or
2DG. The maximum absolute value attained after injection (max.) is given for
each treatment group. See text for levels of statistical significance.
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
Effects of DE on the Responses of Plasma Corticosterone Concentrations to
2DG.
Corticosterone (CORT) responses to vehicle or 2DG are illustrated in Table
2.2 and Figure 2.7. Vehicle injections did not significantly increase plasma
corticosterone concentrations from pre-injection values in any treatment
group on days 1, 3 or 9 (Fig. 21 k). Flowever, 2DG injections significantly
increased plasma corticosterone concentrations on day 3 in EU animals (F
{df 4,35} = 9.61, P< 0.001, Fig. 2.7B). Following 5 days of DE on day 9, preÂ
injection levels of CORT were significantly elevated from those measured at
day 3 (F {df 1,10} = 6.8, P< 0.05, Table 2.2). Flowever, 2DG injections still
significantly increased plasma corticosterone concentrations from preÂ
injection values in DE animals (F {df 4,33} = 3.95, p< 0.01, Fig. 6C). Two-
way ANOVA with repeated measured revealed a significant main effect of
2DG on plasma corticosterone concentrations at the time (60 mins) when
mean maximum value was attained after 2DG injections (F {df 1,10} = 82.12,
p<0.001, Fig. 6C). Finally, there was no significant interaction effect at any
time point. There was also no significant difference between the incremental
increase of CORT in animals receiving 2DG, regardless of hydration state at
any time point after the pre-injection measurement.
50
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
A B
T 3
~ U i
E,
0
w
o
o
.3
O )
~ o
o
o
.Q
0
O )
c
0 3
o
c
200
160
120
80
_ 40
£
c
0
E
0
0
-20
120 0 30 60 90 120
Time (min)
Figure 2.6: The incremental increase in mean (± SEM) plasma glucose
concentrations above pre-injection values.
(0 mins) in animals after injection of vehicle (A), 200 mg/kg 2DG given 1 day
later (B, closed squares), and 200 mg/kg 2DG given after 5 days of
hypertonic saline ingestion (C, closed squares). The change in mean plasma
glucose concentrations after vehicle injection are shown in animals before (B,
open circles) and after 5 days of hypertonic saline ingestion (C, open circles).
DISCUSSION
Our present results demonstrate two points regarding the control of energy
balance in DE-anorexic rats. First, that normal plasma glucose
concentrations are maintained during DE-anorexia, presumably because
51
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metabolism is now biased towards increased glycogenolysis and lipolysis
This observation, taken together with the fact that DE-anorexic and paired-
FR animals have virtually identical endocrine and neuropeptidergicresponses
to negative energy balance (Watts et a i, 1999), shows that DE-anorexic
animals maintain a normal metabolic response to reduced food intake. Of
A B
O )
c
< D
C
o
I n -
C D
■* — »
in
O
g
r
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0 3
c
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_ £ =
O
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E
< 1 )
i—
O
c
400
300
200
100
0
-100
0 30 60 90 120 0 30 60 90 120 30 60 90 120 0
Time (min)
Figure 2.7: The incremental increase in mean (± SEM) plasma
corticosterone concentrations above pre-injection values.
(0 mins) in animals after injection of vehicle A), 200 mg/kg 2DG given 1 day
later (B, closed squares), and 200 mg/kg 2DG given after 5 days of
hypertonic saline ingestion (C, closed squares). The change in mean plasma
corticosterone concentrations after vehicle injection are shown in animals
before (B, open circles) and after 5 days of hypertonic saline ingestion (C,
open circles).
52
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
course, the critical difference between FR and DE-anorexic animals is the
decreased drive to eat in the latter. Second, we show that DE for as little as
three days results in a severe attenuation of the compensatory food intake
that normally occurs after an overnight fast. DE animals also eat less in
response to doses of 2DG that elicit feeding in the same rats before DE, and
have been shown by others to produce eating (Ritter, Ritter, and Cromer,
1999; Singer, and Ritter, 1994). These observations are consistent with a
previous study showing 2DG-induced food intake is attenuated in water-
deprived rats (Watson, and Biderman, 1982). DE animals therefore do not
seek to repair either an actual (from overnight starvation) or a perceived
(from 2DG) caloric deficit until after access to water has been restored.
Similar responses have been reported in rats with activity based anorexia
(Aravich, Stanley, and Doerries, 1995) and interestingly, in humans with
anorexia nervosa. Control patients reported feelings of increased
hungewithin minutes of intravenous administration of 2DG or insulin but
patients with anorexia nervosa reported no increase in their hunger ratings
following these challenges in comparison to infusion of vehicle. This was the
case despite that both the blood glucose and cortisol levels responded to the
challenge in the same manner as did those of the control patients (Nakai,
Kinoshita, Koh, Tsuji, and Tsukada, 1987; Nakai, and Koh, 2001).
At this point DE-anorexic animals reliably begin robust compensatory feeding
within ten minutes from being given back drinking water (Watts, 1999). This
53
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rapid eating response clearly demonstrates that DE animals will eat with
appropriate stimulation, and that the mechanisms responsible for inhibiting
feeding to a variety of stimuli are quickly counteracted by drinking water. The
mechanisms responsible for the reversal of this anorexia are currently
unknown.
2DG-induced glucoprivation rapidly elicits a triad of compensatory motor
responses aimed at mobilizing glucose stores and replenishing energy
supplies: increased secretion of epinephrine to produce hyperglycemia,
glucocorticoid release, and feeding. These complimentary processes are
activated more or less simultaneously and promote glucose delivery to the
brain. However, the fact that under certain circumstances they can be
uncoupled, demonstrates that their control mechanisms are not tightly linked.
For example, phlorizin and alloxan, which inhibit glucose transport and
glucose oxidation respectively, both elicit eating behavior but not
hyperglycemia when injected into the fourth ventricle (Flynn, and Grill, 1985;
Ritter, and Strang, 1982). Similarly, area postrema lesions impair feeding
after 2DG administration but leave intact both hyperglycemic and the
corticosterone secretory response (Edmonds, and Edwards, 1998). In this
regard, we show that DE-anorexic animals retain the ability to mount both a
hyperglycemic and a glucocorticoid secretory response to the same dose of
2DG that fails to stimulate eating. DE therefore specifically targets pathways
54
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
associated with stimulating food intake, while leaving intact those
mechanisms responsible for neuroendocrine and sympathetic
glucometabolism-related motor events.
Repeated daily 2DG administration can impair the feeding response to 2DG
(Sanders, and Ritter, 2000), possibly as a result of a chronic elevation of
circulating glucocorticoids (Davis, Shavers, Davis, and Costa, 1997). In the
present study we have confirmed previous reports that dehydrated animals
show increased plasma corticosterone levels in the morning (Watts, 1992).
However, the mechanisms responsible for suppressing feeding in these
animals are most likely different from those arising after repeated daily 2DG,
and three observations suggest that this suppression is probably not a
consequence of these increased plasma corticosterone concentrations. First,
we show that non-DE control rats receiving two 2DG challenges 7 days apart
show identical feeding responses to each challenge. Second, the morning
elevation in plasma corticosterone levels seen in DE animals remains well
below the peak values attained after 2DG injection in EU animals (present
study; (Watts, 1992). Finally, unlike DE, repeated daily injections of 2DG not
only attenuate the feeding response, they also abolish the 2DG-induced
hyperglycemia (Sanders etal., 2000).
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
The neural mechanisms that control feeding following glucoprivation or
deprivation are not fully understood, but the large body of data implicating
NPY/catecholaminergic neurons in the hindbrain and NPY/agouti-related
protein-containing neurons located in the ARH provides a framework for
discussing our results with regard to the neural substrates of anorexia.
Injections of an anti-dopamine fc-hydroxylase-saporin conjugate (D-SAP) into
the terminal regions of catecholaminergic neurons will specifically destroy
these neurons (Ritter etal., 2001a). Ritter and colleagues have recently
taken advantage of this specificity to show that D-SAP injected into the PVH
blocks both the feeding and corticosterone responses to 2DG, but leaves
intact the hyperglycemic response (Ritter et at, 2001a; Ritter et al., 2001b).
In contrast, D-SAP injected into the spinal cord destroys catecholaminergic
neurons with descending connections and blocks the hyperglycemic
response to 2DG while leaving the feeding and corticosterone response
intact (Ritter et at, 2001a; Ritter et at, 2001b). These data demonstrate that
different subsets of hindbrain catecholaminergic neurons mediate the
behavioral, autonomic, and neuroendocrine components of the glucoprivic
response.
We show that DE-anorexic rats have hyperglycemic and corticosterone
responses to 2DG that are indistinguishable from controls. This
demonstrates that the inhibitory mechanisms present in DE-anorexia rats do
56
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not impact either those ascending and descending catecholamine pathways
that target CRH neuroendocrine neurons and mediate glucocorticoid
responses, or those pre-ganglionic neurons in the spinal cord that mediate
hyperglycemia. In addition, our results do not support the view that
neuroendocrine CRH neurons in the PVH are involved with compensatory
feeding behaviors (Watts etal., 1999); corticosterone secretion remains
viable in DE animals while feeding is markedly impaired. This notion is also
supported by the fact that electrolytic lesions of the PVH do not hinder
glucoprivicfeeding (Ritter etal., 2001a; Ritter etal., 2001b). Furthermore,
DE most likely does not act on NPY neurons in the ARH to inhibit feeding to
DE, as complete ablation of NPY neurons with a saporin-NPY conjugate
does not hinder 2DG feeding (Bugarith etal., 2005). Collectively, these data
suggest that 2DG-induced feeding requires sets of hypothalamic neurons
located outside the PVH and the ARH, and that these systems are potential
targets for DE-generated inhibition.
In conclusion, we have shown previously that the anorexia that develops
after drinking hypertonic saline inhibits spontaneous nocturnal feeding
(Watts, 1999). The present study shows this anorexia also involves an
inhibition of two other types of feeding; compensatory feeding in response to
overnight starvation, and the feeding that usually follows glucoprivation. The
fact that both hyperglycemic and the glucocorticoid responses to 2DG remain
57
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intact in DE-anorexic animals shows that DE specifically targets those
mechanisms that control the motor events of feeding behavior, but not
neuroendocrine or autonomic motor responses.
Our results support the idea that specific alterations to feeding mechanisms
in the ARH are not responsible for DE-anorexia (Watts etal., 1999). In this
regard, several lines of evidence support the idea that a critical component
for the development of DE-anorexia is located within the LHA, particularly its
perifornical part (LHApf). NPY-containing projections from the ARH to the
LHApf are important for stimulating those types of feeding initiated by
changes in the levels of circulating hormones such as leptin (ELIAS et at,
1998b; Elmquist, Elias, and Saper, 1999). Similarly, hindbrain adrenergic and
noradrenergic neurons activated by 2DG co-localize NPY (Sawchenko et al.,
1985) and project to both the PVH and LHA (Elias, Aschkenasi, Lee, Kelly,
Ahima, Bjorbaek, Flier, Saper, and Elmquist, 1999; Swanson, and Hartman,
1975). Furthermore, neurons within the LHA express NPY receptors (Durkin,
Walker, Smith, Gustafson, Gerald, and Branchek, 2000; Gerald, Walker,
Criscione, Gustafson, Batzl-Hartmann, Smith, Vaysse, Durkin, Laz,
Linemeyer, Schaffhauser, Whitebread, Hofbauer, Taber, Branchek, and
Weinshank, 1996) and injections of NPY into the LHApf produce strong
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feeding responses (Stanley et al., 1993). We suggest that DE in some way
inhibits the output of those NPY-containing circuits that normally elicit food
intake in response to caloric deficits. This hypothesis is consistent with
certain other types of anorexia where animals exhibit a suppressed feeding
response to central injections of NPY, have increased NPY gene expression
in ARH, and show increased NPY release in the PVH (Ballinger et al., 2001;
Blanton et al., 2001; Jensen et al., 1998; Rioux etal., 2001). However, unlike
these other models of anorexia, DE-anorexia is rapidly and completely
reversed within minutes simply by restoring access to drinking water, making
it a particularly useful model with which to investigate the neural substrates of
anorexia.
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CHAPTER THREE
DIFFERENTIAL ANATOMICAL PATTERNS OF NEURONAL
ACTIVATION AFTER 2 DEOXY-D-GLUCOSE IN ANOREXIC
ANIMALS
INTRODUCTION
We have shown that DE-anorexic rats limit spontaneous circadian-driven
food intake despite the presence of endocrine and neuropeptidergic profiles
of negative energy balance (Watts et al., 1999). In fact, DE animals share a
virtually identical profile of caloric deficit markers with animals that are food
restricted (FR) to match the intake of the anorexic animals (Watts et al.,
1999). These include body weight loss, undetectable levels of circulating
leptin and insulin, increased concentration of plasma glucocorticoids, and
elevated neuropeptide Y (NPY) gene expression in the arcuate nucleus of
the hypothalamus (ARH). While these endocrine and neural components are
usually considered as being stimulatory for food intake (Baskin et al., 1999;
Dallman et al., 1999; Friedman et al., 1998; Solano et al., 1999), these
processes are less effective in DE-anorexic rats. In fact, not only does DE-
anorexia reduce spontaneous food intake but it also limits food intake
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following overnight starvation and 2-deoxy-D-glucose (2DG)-induced
glucoprivation (Salter and Watts, 2003; Watts and Salter, 2004).
2DG is a competitive glucose analog that inhibits intracellular glucose
utilization. The resulting cytoglucopenia activates compensatory motor
responses including hyperglycemia driven by sympathetic activation of
adrenomedullary epinephrine secretion (Scheurink et al., 1993);
corticosterone secretion from the adrenal cortex (Weidenfeld et al., 1994),
and food intake (Ritter et al., 1999). These three motor events serve to
mobilize glucose stores and replenish energy supplies, and although they
usually are activated synchronously they can be uncoupled under certain
circumstances. Although DE-anorexic rats mount normal glucoregulatory
hyperglycemic and glucocorticoid secretory responses to 2DG they do not
demonstrate glucoprivic feeding responses (Salter and Watts, 2003).
The targeted suppression of food intake suggests that the output of feeding
stimulatory pathways are inhibited by DE. In the current study, we use 2DG
as a probe to investigate the underlying neural circuitry of anorexia. To do
this we have performed a detailed neuroanatomical comparison of Fos-
immunoreactivity (ir) response patterns in DE and euhydrated (EU) rats given
200 mg/kg 2DG or vehicle. The expression of Fos protein is well-established
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as a functional marker of activated neurons (Hoffman, and Lyo, 2002). Fos
protein increases in the nucleus of activated neurons within 30-45 minutes
following the presentation of a stimulus and has a half life of approximately
two hours (Morgan, and Curran, 1989). We hypothesize that by comparing
the Fos-ir response patterns following 2DG in EU animals (where 2DG
stimulates feeding) with DE-animals (where the feeding response to 2DG is
markedly attenuated) we can identify areas where DE pathways might act to
inhibit feeding.
MATERIALS AND METHODS
ANIMALS AND PROCEDURES
Adult male Sprague Dawley rats weighing 240-260 g were obtained from
Harlan laboratories and singly housed in suspended Plexiglas cages with
sanitized wood chips. They were maintained in a temperature-controlled
room on a 12:12 h light-dark schedule with lights on at 0600. Rats were
provided with continuous access to food (Teklad rodent diet 8604) and water
throughout the experiment except where stated. In some animals, drinking
water was replaced with 2.5% saline solution for 5 days. We have previously
shown that 5 days of drinking hypertonic saline increases plasma osmolality
by approximately 6% (Watts etal., 1995c; Watts e t a i, 1999). Body weights
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and nocturnal food intake were measured daily throughout the experiment.
All procedures have been approved by the local institutional animal care and
use committee.
Four groups of animals were maintained on ad lib food intake and either
water or 2.5% saline for five days. On the morning of the fifth day, food was
removed from all cages. Two groups of dehydrated animals (DE) or
euhydrated (EU) animals were given an subscapular injection of either 2DG
(200mg/kg, DE-2DG, EU-2DG) or an equal volumetric injection of
physiological saline (DE-VEH, EU-VEH). This dose of 2DG was chosen
because it generated a sub-maximal set of responses in control animals
(Salter et al., 2001).
HISTOLOGY AND FOS IMMUNOCYTOCHEMISTRY
Two hours after injection, animals were anesthetized with halothane
anesthesia and rapidly decapitated. Brains were removed from the skull and
immersion-fixed for three days on ice cold PBS 4% paraformaldehyde and
then frozen on dry-ice and stored at -70°C. We have previously shown that
this fixation method is compatible with the preservation of Fos-ir (Khan, and
Watts, 2004; Sanchez-Watts, Kay-Nishiyama, and Watts, 2000). Two sets of
five series of 1-in-5, 30 fim thick frozen coronal sections were cut through the
brain. The first ran from the level of the subfornical organ to the caudal part
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of the arcuate nucleus (ARH); the second ran from the parabrachial nucleus
through the caudal hindbrain. Sections were stained for Fos im mu no reactivity
using previously published methods (Kelly etal., 1996). Briefly, after rinsing
with potassium phosphate buffered saline (KPBS), sections were incubated
in Ab-5 anti-c-fos antibody (1:40K, Oncogene Science, Cambridge, MA) for
48 hours, with 3% normal goat serum and 0.2% Triton X-100. Sections were
rinsed and incubated for 2 hours with biotinylated 2° antibody, followed by
incubation with Vectastain Elite ABC reagent (Vector Labs, Burlingame CA).
Specific antibody staining was color-detected with a 1 mg/ml solution of
daminobenzidine (DAB, 3,3 diaminobenzidine tetrahydrochloride) with 0.3%
hydrogen peroxide. Adjacent sections were Nissl stained. All sections were
mounted, cover slipped and examined under the microscope.
IMAGE ANALYSIS
Brain areas to be analyzed were selected based on their known involvement
in feeding- related pathways. Anatomically defined regions of the
hypothalamus and hindbrain were then located on the stained sections using
landmarks identified using the Swanson Rat Atlas (Swanson, 1998) and
adjacent Nissl-stained sections. Images of each field were printed and Fos-ir
labeled cells counted manually and verified by microscopic examination of
the original slide and its corresponding nissl-stained section. Labeled cells
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on each side of all regions were counted independently of the other, without
knowledge of the experimental group. Because the number of sections
needed to span the entire region of interest differed from animal to animal
and from side to side, the cell counts were expressed as labeled cells-per-
section (total number of cells per region/total number of sections per region).
The mean number of cells-per-section was then compared across groups.
ANOVA was performed on groups with statistical significance established at
P<0.05.
Paraventricular Nucleus of the Hypothalamus
Labeled cells were counted in a rostrocaudal series that ranged from the
rostral point of the lateral zone of the posterior magnocellular part through
the forniceal component (level 26 to level 28 of the Swanson Atlas). Cell
counts were further divided by close examination with adjacent Nissl sections
into posterior magnocellular component (lateral zone), medial parvicellular
part (dorsal zone), and lateral parvicellular and forniceal components.
Lateral Hypothalamic Area
Labeled cells were counted from the caudal extent of the forniceal
component of the paraventricular nucleus (level 27) to the rostral extent of
the posterior part of the dorsomedial nucleus (level 30). Neurons were
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counted ventral to the dorsal limit of the third ventricle, medial from the
internal capsule, lateral to the medial margin of the fornix and lateral border
of the arcuate, and dorsal to the supraoptic commissure/optic tract. Cells in
the PVH, anterior hypothalamic nucleus, ventromedial nucleus, and
dorsomedial nucleus were excluded from the LHA counts. These LHA areas
were chosen to include the perifornical area of the hypothalamus as well as
the areas where melanin-stimulating hormone and orexin/hypocretin-
producing neurons have been reported to be found, as they have been
implicated in feeding behavior (Broberger, De Lecea, Sutcliffe, and Hokfelt,
1998a; Stanley etal., 1993).
Arcuate Nucleus of the Hypothalamus
Fos-ir labeled cell counts began for the ARH at the rostral edge of the
ventromedial nucleus (level 26) through to the rostral extent of the posterior
dorsomedial nucleus (level 29). Any labeled neurons in the median eminence
were excluded.
Parabrachial Nucleus
Labeled cells were counted beginning where the dorsal tegmental nucleus is
dorsal to the medial longitudinal fasiculus (level 48) through to the area
where the superior cerebellar peduncle meets the ventral spinocerebellar
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tract (level 51). In the PBN region Fos-ir neurons were counted in the lateral
component (central, external, and ventral lateral part of the PBN).
Area Postrema
Cells containing Fos-ir were counted throughout the entire extent of the area
postrema (level 69-70). Any labeled neurons seen in the nucleus gracilus
and commissural part of the nucleus of the solitary tract were excluded.
Nucleus of the Solitary Tract/Dorsal Motor Complex
Cell counts began at the rostral pole of the area postrema (level 67) and
continued through to the caudal limit of the area postrema (level 70). Cells
were counted in the medial, commissural, gelatinous, and central part of the
NTS, and the dorsal motor complex dorsal to the hypoglossal nucleus. The
neurons in the area postrema were counted separately.
RESULTS
Figures 3.1 &3.2 illustrate the response of Fos immunoreactivity in
euhydrated animals after either 2DG or vehicle injection (EU-2DG, EU-VEH),
and dehydrated animals after either 2DG or vehicle injection (DE-2DG, DE-
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VEH). Figures 3.3 and 3.4 show graphically the neuronal areas where Fos-ir
increased after 2DG in DE (Fig 3.3), and those without an increased Fos
response to 2DG (Fig 3.4).
Vehicle 2-DG DE DE-2DG
PVH
mpd & pmI
ARH
PBN
AP/NTS
G E F H
... scp
p N
ill
Figure 3.1: Fos immunoreactivity after 2DG administration in euhydrated and
dehydrated animals.
Brightfield photomicrographs of representative coronal sections showing Fos
immunoreactivity in the paraventricular nucleus (PVH, A-D), arcuate nucleus
(ARH, E-H), parabrachial nucleus (PBN, l-L), and area postrema, nucleus of
the solitary tract (AP/NTS, M-P) of animals in the four treatment groups
sacrificed two hours after injection of vehicle in euhydrated animals (A,E,I,M)
and DE animals (C,G,K,0) and after injection of 2DG in euhydrated animals
(B,F,J,N) and DE animals (D,H,L,P). 3v, third ventricle; scp, superior
cerebellar peduncle, sol, solitary tract. PVHmpd, dorsal aspect of the medial
parvicellular part of the PVH and PVHpml, posterior magnocellular part of
PVH.
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2-DG
LHApf
Vehicle 2-D G DE D E -2D G
. ■- . ••
P .. '
<3, .. • h •
.
’ • F •
: : ' r â– . â– â–
V y : ; â– -/-
. • - • •
V F . * * ‘
*
Figure 3.2: : Fos immunoreactivity after 2DG administration in euhydrated
and dehydrated animals.
Brightfield photomicrographs of representative coronal sections showing Fos
immunoreactivity in the lateral parvicellular part of the paraventricular
nucleus (PVHIp, A-D) and the perifornical part of the lateral hypothalamic
area (LHA, E-H) of animals in the four treatment groups sacrificed two hours
after injection of vehicle in euhydrated animals (A,E) and dehydrated
animals (C,G) and after injection of 2DG in euhydrated animals (B,F) and DE
animals (D,H,). F, fornix.
Hypothalamus
Figure 3.1 and 3.2 illustrate the Fos-ir response patterns in the medial (Fig
3.1) and lateral parvicellular (Fig 3.2), and the posterior magnocellular
components of the PVH (Fig 3.1). When Fos-ir neurons were counted in the
medial parvicellular component of the PVH (PVHmpd), 5 days of drinking
hypertonic saline did not significantly raise Fos activation levels over control
69
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animals alone, whereas both 2DG alone and 2DG in DE significantly
elevated Fos in this nucleus (Fig 3.3). However, 2DG given to DE animals
significantly reduced Fos-ir from that of animals given 2DG alone (Fig 3.3,
P<0.05).
There was no significant increase in Fos-ir in the lateral parvicellular
component of the PVH (PVHIp ) after 2DG administration. Fos in the PVHIp
did significantly increase over control animals after DE, with or without 2DG
administration (Fig 3.4). Particularly, there was a noticeable increase of Fos-
ir in the periforniceal component of the lateral wing of the PVH. Five days of
drinking hypertonic saline resulted in increased neuronal activation in the
magnocellular neurons (PVHpml) compared to controls (Fig 3.4).
Administration of 2DG also increased the number of fos-ir positive neurons
over that seen in control animals. When 2DG was administered to DE
animals, there were more neurons showing Fos activation than in control
animals in the PVHpml (Fig 3.1).
Analysis of Fos immunoreactivity in the LHA (Fig 3.4) demonstrated an
increase in activation over control animals after 2DG and after 5 days of
drinking hypertonic saline (Fig 3.4). In contrast, when DE animals were
administered 2DG Fos-ir, although still significantly higher than control
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animals, the number of activated neurons was not significantly higher than
that seen when the stimuli were provided independently of one another (Fig
3.4).
Either 2DG administration or DE alone increased Fos-ir n the ARH
(Fig 3.1 &3.3) above that seen in control animals. 2DG administered to DE
animals further increased Fos-ir above that seen in control, 2DG and DE
animals (Fig 3.3).
Parabrachial Nucleus
The number of neurons in the lateral portion of the parabrachial nucleus (Fig
3.1) with Fos-ir nuclei was significantly higher than control animals after
either 2DG administration or DE. When 2DG was administered to DE
animals, there were more neurons showing Fos-ir than control animals,
which was more than when the stimuli were given independently (Fig. 3.3).
Nucleus of the Solitary Tract, Dorsal Motor Nucleus of the Vagus, and Area
Postrema
5 days of drinking hypertonic saline significantly elevated the number of Fos-
ir nuclei in the NTS and DMX over that seen in control animals (Fig 3.1 &
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3.3). Likewise, 2DG administration alone and 2DG administered following DE
significantly elevated Fos-ir above control amounts at these sites.
250
200 â–
C D
0
1 150
C D
.Q
T O
C D
I 100
3
z
50
*
X
â– *#
NS â– *
II ooii
AP NTSm/DMX PBI PVHmpd ARC
Figure 3.3: Mean (±SEM) number of Fos positive neurons after vehicle or 2-
DG administration in EU or DE animals.
Shown are control vehicle-injected animals (white bars), 2DG-injected (light
gray bars), 5 days of DE (dark gray bars), and 2DG administered after 5
days of DE (black bars). Symbols above bars represent significant difference;
different from vehicle-injected controls (*), different from other indicated
group (**), difference from vehicle, 2DG, and DE (#), and not significant from
vehicle injected control (NS). See text for detailed statistical significance.
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Counts of Fos-ir cells in the AP (Fig 3.3) were similar to those seen in the
NTS. When 2DG was administered to DE animals, more nuclei showed Fos-
ir than in control animals, and there were more Fos-ir nuclei than when the
NS
w
1 5
O
T J
C D
.O
c n
C D
-Q
£
3
300i
250-
200-
150-
100-
LHA PVHpml PVHIp
Figure 3.4: Mean (±SEM) number of Fos-positive neurons after vehicle or 2-
DG administration in EU or DE animals.
Shown are control vehicle-injected animals (white bars), 2DG-injected (light
gray bars), 5 days of DE (dark gray bars), and 2DG administered after 5
days of DE (black bars). Symbols above bars represent significant difference;
different from vehicle-injected controls (*), not statistically significant from
indicated groups (NS). See text for detailed statistical significance.
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animal received either 2DG or DE independently. Each of the test conditions
resulted in a significant increase over the number of Fos positive neurons
counted in control animals (Fig 3.3).
DISCUSSION
We have previously shown that DE significantly attenuates feeding after 2DG
administration while leaving other glucoregulatory responses intact (Salter et
at., 2001, see chapter 2). These findings suggest that some component of
the neural network responsible for activating eating is preferentially inhibited
by DE compared to those networks that stimulate corticosterone or
adrenaline secretion. The current study is designed to identify potential
neural substrates of this inhibition by comparing patterns of Fos-ir stimulated
by 2DG in control and DE-anorexic animals. We reason that regions with few
or attenuated numbers of Fos-positive neurons in DE animals following 2DG
administration are good candidates for being associated with this inhibition.
Although quantifying Fos-ir is commonly used to identify activated neurons, it
is not without limitations. Increased Fos expression involves either increasing
intracellular calcium or cyclic AMP and although these events can result from
depolarization, increases in both Fos expression and a neuron’s firing rate
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are dissociable (Berretta, Robertson, and Graybiel, 1992; Luckman, Dyball,
and Leng, 1994). Therefore, increased Fos-ir is best considered a measure
of the afferent input (integrated receptor activation) to a particular neuron,
rather than its output (firing rate). Any reduction or elimination of 2DG-
induced Fos activation by DE we see in this study may indicate where 2DG
related-afferent activity is altered by DE.
With this in mind, our results demonstrate three points. First, when
administered separately DE and 2DG each elevate Fos-ir in several key
areas considered important for neuroendocrine regulation and for controlling
ingestive behaviors. Second, when 2DG is administered to DE-anorexic
animals the number of neurons showing Fos activation in most of these
same areas is increased in a simple additive manner, suggesting that they
are not integral parts of the circuit that inhibits feeding. The AP is an
exception here, in that the numbers of Fos-ir neurons was greater when the
stimuli were combined than when they were administered separately. Third,
some of these same cell groups, notably the LFIA and PVHIp, showed
increased Fos-ir to one or both challenges when given separately, but failed
to show further increases when given together. Collectively, these
observations suggest that the neurons in the LHA, PVHIp, and AP can, at
least in part, mediate DE-anorexia.
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The ARH is traditionally considered to regulate the behavioral, endocrine,
and autonomic motor responses to altered energy status, particularly those
signaled by changes in circulating leptin and insulin (Elmquist, Maratos-Flier,
Saper, and Flier, 1998b). A hypocaloric state is characterized by sharply
lower circulating levels of insulin and leptin, which reduce POMC gene
expression and increase NPY and AGRP gene expression in ARH neurons
(Sahu, 1998b; Sahu, 1998a; Schwartz, Seeley, Woods, Weigle, Campfield,
Burn, and Baskin, 1997). Chronically DE rats are also severely hypocaloric
(Watts et a!., 1999), and show the same changes in ARH gene expression as
food restricted animals (O'Shea etal., 1995; Watts et a!., 1999).
In agreement with previous studies (Minami etal., 1995) we found that 2DG
administration increased Fos-ir in ARH cells of EU animals. Similarly, like
previous research, the hypertonic stimuli of DE also increased Fos-ir in the
ARH (Solano-Flores, Rosas-Arellano, and Ciriello, 1993). When 2DG was
administered to DE animals, the numbers of Fos-ir neurons were
approximately double those seen when 2DG or DE were administered
separately. One explanation of this observation is that there are two distinct
populations of Fos-responsive neurons in the ARH: one that responds to
2DG, and one that responds to DE. Although it appeared that more Fos -ir
neurons were present in the medial component of the ARH after 2DG than
after DE, the neuropeptidergic content of the activated neurons in response
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to each stimuli is not known. Regardless, DE does not appear to inhibit the
Fos response of the 2DG-sensitive population. This conclusion is consistent
with our previous findings showing that ARH neurons in DE animals remain
capable of responding appropriately to signals of diminished energy status,
at least in terms of gene expression (Watts et a/., 1999), also see Chapter 5.
Furthermore, the fact that glucoprivic feeding is normal after complete
chemical ablation of ARH NPY neurons (Bugarith etal., 2005) supports the
notion that the inhibition of 2DG feeding by DE is not initiated at the level of
the ARH. Rather, our more recent findings suggest that one reason that DE-
animals remain anorexic is the significantly reduced sensitivity of some of
their NPY target sites, particularly the PVH and LHA (Salter and Watts,
2003a; Salter and Watts, 2004; Watts and Salter, 2004),
Caudal hindbrain structures located in the dorsomedial medulla, such as the
medial components of the NTS and the AP have long been posited as crucial
components of the glucoprivic response after administration of glucose
analogues (Ritter, Slusser, and Stone, 1981). Anatomically, the AP is a
superior candidate for housing neural substrates capable of glucoreception.
Fenestrated capillaries allow easy access of blood-borne signals to the AP.
Furthermore, the AP is linked bidirectionally with the NTS (Hay, and Bishop,
1991) a site of convergent chemosensitive vagal fibers and cell bodies that
send neuronal efferents to the ventral lateral medulla (Chan and Sawchenko,
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1995), the parabrachial area (Fulwiler and Saper, 1984) and the PVH
(Sawchenko etal., 1982). Thus, in agreement with previously published
results (Ritter etal., 1998), we found that peripheral administration of 2DG
increased the numbers of Fos-ir neurons in the NTS, DMX, AP, and PB.
However, in terms of understanding the circuitry that mediates anorexia
during DE, it is important to note that DE did not reduce the numbers of Fos-
ir neurons following 2DG in the NTS, DMX, or PB. Indeed, 2DG
administration in DE animals increased the number of of Fos-ir neurons in
these three areas to values that were close to the sum of those seen
following separate administration. This fact, along with the similar
observation in the ARH, suggests that DE and 2DG each target distinct
populations of neurons in the NTS, DMX, and PB.
Interestingly, the simple additive pattern we saw in other hindbrain regions
was not apparent in the AP. Although 2DG or DE alone elicited Fos
activation, when 2DG was given to DE animals there were significantly more
Fos-ir neurons than when each stimulus was presented separately. This
observation is consistent with the existence of a third population of AP
neurons that is only activated by a combination of DE and 2DG, and this
population may be one that can inhibit the pathways responsible for
stimulating food intake in response to 2DG. However, this notion is not
parsimonious with research showing that the complete removal of the AP
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selectively impairs the feeding response to 2DG (Edmonds etal., 1998). It is
unlikely that an increase in neural activation in the AP elicits the same
behavioral response as does the removal of critical neural substrates.
Fos-ir was substantially increased following DE in the PVHIp. However,
unlike the ARH and some hindbrain structures, we found the PVHIp
responded differently from the other cell groups we examined in that DE
alone increased the numbers of Fos-ir neurons in the PVHIp, while 2DG had
no effect in either EU- or DE-animals. The PVH contains sets of parvicellular
neurons that project to the midbrain, hindbrain and spinal cord and are
heavily implicated in autonomic and behavioral control (Hallbeck, and
Blomqvist, 1999; Hallbeck, Larhammar, and Blomqvist, 2001; Swanson, and
Kuypers, 1980). These results, when viewed in light of several other lines of
evidence, support the notion that the parvicellular neurons within the PVHIp
are chronically activated by DE (Stocker, Cunningham, and Toney, 2004)
and that these neurons are potential contributors to DE-generated inhibition
of feeding.
We found two other regions containing neuroendocrine motor neurons also
did not show an additive response as seen in the ARH. In the PVHpml, which
contains AVP and OT magnocellular neurons, we found significant numbers
of Fos-ir cells following DE. Although 2DG administration increased Fos-ir in
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these magnocellular neurons, the number of Fos positive neurons was less
than after DE alone. When DE animals were given 2DG, there was little
further change in the Fos-ir cells, most likely because these neurons were
already activated by the five days of DE. Why 2DG alone increased
activation in these magnocellular neurons is unclear, although it may result
from osmotic stimuli associated with the sustained hyperglycemia that occurs
after 2DG administration (Iwasaki, Kondo, Murase, Hasegawa, and Oiso,
1996; Salter et al., 2001).
In contrast to magnocellular neurons, we found that 2DG substantially
increased the numbers of Fos-ir neurons in the PVHmpd, which is consistent
with other reports in the literature (Ritter et al., 1998). This PVH compartment
contains the neuroendocrine CRH neurons responsible for increased
glucocorticoid secretion in response to 2DG (Ritter, Watts, Dinh, Sanchez-
Watts, and Pedrow, 2003). Interestingly, when 2DG was given to DE
animals, the Fos-ir response was decreased from that seen in EU animals.
This finding is consistent with the diminished responsiveness of these
neurons in DE animals to some stressors (Watts, 1996; Watts and Sanchez-
Watts, 2002)
Although the Fos-ir response profile of the neurons in these two
neuroendocrine compartments is consistent with one that might generate
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anorexia (ie. the Fos response to 2DG is attenuated by DE), four
observations make it unlikely that these neurons can be directly implicated in
this manner. First, these neurons are neuroendocrine and projections to
regions other than to the neurohypophysis have not been reported. Second,
DE-anorexia does not result from either elevated corticosterone or reduced
CRH mRNA levels in the PVHmp because adrenalectomy abolishes the
reduction in CRH mRNA following DE but not the development of anorexia
(Watts and Sanchez-Watts, 1995d; Watts etal., 1999). Third, 2DG-
stimulated corticosterone secretion remains viable, while feeding is
attenuated in DE animals (Salter and Watts, 2003b), see also Chapter Two .
Finally, animals continue to eat to 2DG even after electrolytic destruction of
the PVH (Calingasan et al., 1992; Shor-Posner, Azar, Insinga, and
Leibowitz, 1985).
The third pattern of Fos response we observed to 2DG in DE animals was
the one seen in the LHA . Although 2DG increased Fos activation in the
LHA this increase was not elevated further when 2DG was administered to
DE animals despite the fact that DE alone elicited a robust number of Fos-ir
neurons. Animals with excitotoxic LHA lesions continue to regulate ingestive
behaviors appropriately after adulteration of their water or food supply, and
they can still compensate appropriately to food or water deprivation (Clark,
Clark, and Winn, 1990; Clark, Clark, Bartle, and Winn, 1991). Critically
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however, LHA lesioned animals do not eat after a glucoprivic challenge, nor
do they drink appropriately after an injection of hypertonic saline (Clark,
Clark, Warne, Rugg, Lightman, and Winn, 1991; Winn, 1995). LHA lesions
therefore do not hinder compensatory responses when homeostatic
challenges are perceived both by internal metabolic alterations and external
sensory information, as in the case of absent food or water sources.
Alternatively, LHA lesioned animals cannot compensate appropriately to
homeostatic challenges when the deficit is signaled entirely through internal
metabolic alterations.
Cumulatively, these data suggest that some neurons in the LHA can
integrate information from a variety of interosensory and exterosensory
modalities and then initiate appropriate behavioral responses, perhaps
through their extensive projections to the telencephalon and hindbrain (Winn,
1995). This model is consistent with evidence showing the LHApf is a target
of rostral afferents that are sensitive to plasma osmolality (Kelly et al., 1996)
as well as a target for NPY-containing projections from the ARH and the
hindbrain that are important for stimulating feeding after energy deficit and
2DG administration (Bugarith et al., 2005; Elmquist, Ahima, Elias, Flier, and
Saper, 1998a). Thus, the LHApf may house neuronal populations that can
integrate information about hydration state and energy status. Our data
suggest that the decreased Fos-ir response to 2DG in the LHA of DE animals
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may reflect a DE-related alteration of this integration. Thus, the information
coded in NPY-containing circuits that normally elicit food intake in response
to caloric deficits is no longer effective in DE. This notion is further supported
by our research demonstrating the LHA is progressively desensitized to the
feeding stimulatory effects of exogenous NPY during DE (Salter and Watts,
2003a), see also chapter four.
Potential mediators of this inhibition are CRH neurons in the fusiform nucleus
of the bed nucleus of the stria terminalis (BSTfus) and the subset of neurons
within the LHA that contain both CRH and NT. CRH and NT gene expression
both increase in these neurons as soon as these animals begin to drink
hypertonic saline, but before they develop anorexia (Watts, 1999; Watts et
al., 1995a; Watts etal., 1999). Furthermore, not only does the level of CRH
mRNA in the LHA correlate with the strength of the anorexia but CRH mRNA
levels revert to normal within 24 h after anorexia is reversed by drinking
water (Watts et al., 1999). Both CRH and NT are well-documented anorexic
agents; intraventricular injections of CRH inhibit deprivation-induced eating
and NPY-induced feeding, while central injections of NT also reduce food
intake (Levine, Kneip, Grace, and Morley, 1983; Morley, 1987).
DE, as a result of chronic ingestion of hypertonic salt water, and
glucoprivation, as a result of peripheral 2DG administration, both result in
elevated Fos activation in several areas implicated in the control of ingestive
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behaviors. Further, the Fos activation is further increased in most of these
areas when 2DG is administered to DE-anorexic animals including the ARH,
an area that is commonly recognized for its involvement with compensatory
food intake after food deprivation. Therefore, our data support the notion that
the ARH is not responsible for the suppression of food intake seen in DE-
anorexic animals given 2DG. Rather, we suggest that two regions not
demonstrating an increase in Fos activation after 2DG, the LHA and the
PVHIp are the areas that may be involved in the suppression of food intake in
DE-anorexia.
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CHAPTER FOUR
ATTENUATION OF FEEDING RESPONSE AFTER
HYPOTHALAMIC INJECTION OF NEUROPEPTIDE Y IS SITE-
SPECIFIC IN ANOREXIA
INTRODUCTION
Energy balance is maintained by regulating energy intake, use and
allocation. Typically, negative energy balance activates a strong and
powerful metabolic response to preserve lean body mass while activating
central circuits that promote energy intake and storage. The hyperphagia
that normally follows caloric deficit restores energy reserves with a degree of
relative precision (Keesey etal., 1997; Rothwell et al., 1979; Weigle, 1994).
When anorexia is present, however, the drive to increase energy intake in
the face of negative energy balance is reduced or eliminated. Although
anorexia complicates the prognosis of many clinical conditions, the
mechanisms by which appetite and food intake are inhibited in anorexia are
still not well understood.
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Critical to the initiation of hunger and subsequent food intake after a
hypocaloric challenge are several peripheral mediators of peripheral
metabolism. Circulating insulin and leptin levels fall after a missed meal,
while glucocorticoid levels increase (Dallman et al., 1999). Together, these
stimuli provide afferent signals to the hypothalamus, which in turn increases
the drive to eat in part through activation of NPY neurons in the basomedial
forebrain, namely the arcuate nucleus (ARH). These neurons project to the
perifornical lateral hypothalamic area (LHApf) and the paraventricular
nucleus of the hypothalamus (PVH), areas that, when injected with NPY,
elicit robust eating (Broberger etal., 1999; Elias, Kelly, Lee, Ahima, Drucker,
Saper, and Elmquist, 2000; Stanley et al., 1984; Stanley etal., 1993).
Additionally, manipulations that result in an increased food intake, such as
food deprivation or diabetes, increase NPY synthesis and release, tissue
content and gene expression within this circuit (Beck et al., 1990; Kalra et
al., 1991; Sahu et al., 1988; Williams et al., 1989). Thus, activation of the
ARH-PVH-LHA neural network is critical for refeeding after energy deficit.
Persistence of anorexia despite multiple markers of negative energy balance
suggests a failure of NPY driven mechanisms to stimulate adaptive feeding
responses.
NPY gene expression is increased in the ARH in DE-anorexic rats similar to
that seen in starved animals. In addition, DE inhibits compensatory feeding
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responses to overnight starvation and 2 deoxy-D-glucose (Salter and Watts,
2003b) see also chapter two, two metabolic challenges in which eating
behaviors are stimulated through NPY-mediated mechanisms from separate
afferent systems (Bugarith etal., 2005; Elmquist etal., 1998a), suggesting
the output of NPY pathways is inhibited in DE. The present study was
designed to investigate the specificity of this inhibition. Testing the sensitivity
of ARH-NPY terminal sites, specifically the PVH and LHApf, to exogenous
NPY administration should provide key information as to the nature of the
inhibition of feeding in DE.
MATERIALS AND METHODS
ANIMALS AND PROCEDURES
Adult male Sprague-Dawley rats (240-260 g) obtained from Harlan
laboratories were individually housed in suspended Plexiglas cages with
sanitized wood chips. They were maintained in a temperature-controlled
room on a 12:12-h light-dark schedule with lights on at 0600. Rats were
provided continuous access to food (Teklad rodent diet 8604) and water
throughout the experiment, except where stated. In some animals, drinking
water was replaced with during 2.5% saline solution for up to five days. Body
weights and food intake were measured through out the experiment. All
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procedures have been approved by the local institutional animal care and
use committee.
SURGICAL PROCEDURES
Rats were handled daily for approximately 5 days before any surgical
intervention and daily thereafter. Guide cannulas were fabricated from 26
gauge stainless steel tubing and occluded with a removable stainless steel
obturator. For stereotaxic implantation of the guide cannula, rats were
anesthesized with an injection (100pJ/kg i.m.) of ketamine/
xylazine/acepromazine cocktail solution (5 ml ketamine HCI.100 mg/ml; 2.5
ml xylazine, 20mg/ml; 1 ml acepromazine, 10 mg/ml; 1.5 ml 0.9% saline
solution). The skull was exposed and trephined at the implantation site using
flat-skull coordinates of 1.8 mm caudal to Bregma, 0.2 mm off midline and
6.5 mm ventral to the dura for injections aimed to the central portion of the
PVH. For injections to the LHApf stereotaxic coordinates were 2.0 mm
caudal to Bregma, 1.5 mm off midline and 6.9 ventral to dura. Guide cannula
were lowered to the desired site and anchored to the skull with dental acrylic
and stainless steel screws. After the skin incision was sutured, rats were
given an injection of analgesic (Banamine HCI, 2.5 mg/ml, 100pil/kg i.m.),
allowed to recover from anesthesia and were then returned to a clean home
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cage. Behavioral testing began 7-9 days post-surgery, when body weight and
food intake had returned to pre-surgical values.
INJECTIONS
All injections were given between 0900-1000 h in equal volumetric doses of
300 r|l over 60-120 seconds. The obturator was removed and replaced with
a 33 gauge stainless steel injection cannula attached by polyethylene tubing
to a 1 pi Hamilton syringe. The injection cannula extended 1.5mm past the tip
of the guide cannula to facilitate the entry of either NPY or aCSF into the
parenchyma. During injections, the movement of a small bubble in the
calibrated infusion line was used to verify injectate delivery. In addition, the
system was tested before and after each injection to ensure it was free of
leaks or plugs.
FEEDING TESTS
Immediately following the injectate delivery, the injection cannula was
removed, the obturator was replaced into the guide cannula, and the rat was
returned to its home cage for feeding tests with water or 2.5% saline to drink
and a pre-measured amount of rat chow. Food consumption was measured
to the nearest 0.1 g by weighing food and crumbs remaining in cages each
hour for four hours total. Feeding latency was also measured.
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EXPERIMENTAL TIMELINE
The experimental timeline is illustrated in figure 2.1. Rats were injected with
artificial CSF (aCSF) to obtain baseline measurements. Two days later,
animals were injected with NPY (Peninsula Laboratories) dissolved in aCSF.
Equal numbers of rats were tested with NPY first, to ensure a counterÂ
balanced design. Rats were then weight-matched and placed into groups.
After a rest period of 2 days, drinking water was replaced with 2.5% saline for
DE groups (DE-NPY and DE-VEFI). Euhydrated (EU-NPY) weight-matched
control rats remained on drinking water. Food intake and weights were
monitored. After three days of DE, when nocturnal intake was approximately
50% of pre-DE intake, rats were again tested for their feeding response to
injection of either vehicle or the same dose of NPY given previously. We
have previously shown that plasma osmolarity is increased by - 6% at this
stage of DE (Watts et ai, 1995a). Each EU animal was also given a second
dose of NPY directly following that of its weight-matched (pre-dehydration)
DE partner. Food intake was measured for four hours, as before. After the
four-hour food test, DE rats’ water was replaced and food intake was
monitored for another hour.
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Food intake meaured j 1-4h 1-4h 1-4h 1h
7-9 days
2 days 2 days
Cannulate vehicle NPY
or NPY or vehicle
(counter balanced)
| 3 days | 4 hours |
Dehydration
(DE groups)
or ad lib water
(EU groups)
NPY
(EU and DE)
or vehicle
(DE)
Water returned
to DE groups
Figure 4.1: Timeline for NPY experiments. Animals were given the first
injection of VEH or NPY 7-9 days following surgery. Each animal then
received an injection of NPY or VEH two days later in a counterbalanced
design. Animals were placed into groups, two groups being DE and one
group continuing on ad lib water intake. At the end of the DE period (3-5
days) all animals were tested for their feeding response to either NPY or
VEH and EU animals were tested for their response to NPY. DE animals
were given water back at the conclusion of the four hour test.
HISTOLOGICAL ANALYSIS
The following day after behavioral testing, animals were injected through the
guide cannula with 300 rjl of a 1% solution of bromophenol blue (Feldberg,
and Fleishhauer, 1959). After ten minutes, animals were anesthetized with
halothane and rapidly decapitated. Brains were removed and immersion-
fixed for three days in ice cold PBS 4% paraformaldehyde. Two adjacent
single series of 40 mm frozen coronal sections were cut through the injection
site. For observation of the bromophenol blue dye injection sites, one series
was mounted onto slides and immediately cover slipped while the second
series was mounted and stained with thionin for clear observation of
structures adjacent to injection sites. Injection sites were localized by using
landmarks identified with the Swanson Brain Atlas (Swanson, 1998). The
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injection site was mapped microscopically using the areas of dye and tissue
damage as markers of the injection. NPY targeted areas were considered to
be areas along the injection tract between the most ventral point of visibly
damaged tissue and the ventral edge of the guide cannula. The assignment
of animals into PVH or LHA groups was based on the histologically derived
location of their injection site. There were 9 animals with injection sites
distant to either of these anatomical loci and they were associated with
limited food intake after NPY injections so were discarded from further
analysis. Injection sites are graphically illustrated in Fig 4.2.
STATISTICAL ANALYSIS
Data are expressed as means ± SEM. Differences in NPY feeding before
and after DE were analyzed using a one-way ANOVA with repeated
measures. Results from EU and DE animals were compared using a 2x2
repeated measure (RM) ANOVA. Bonferroni's pairwise multiple comparisons
test was used to measure individual differences, with critical level for
significance set at p < 0.05.
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28
27
Figure 4.2: Mapping of injection sites. Areas of PVH-targeted injections
are shown in blue in series 26 and 27, areas of LHApf-targeted injections
are shown in darker blue in series 27 and 28. There were 9 animals with low
intake (average four hour intake after injection 3.6 ±0.6 g) which are outside
target areas and are marked by red stars. Maps are illustrated on sections
from Swanson’s rat brain atlas (Swanson, 1998).
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RESULTS
Experiment 1- injection of NPY to the PVH after 3 days of DE
After injection of either 0.5 or 1.0 |jg of NPY, animals ate significant amounts
of food (p< 0.001, figure 4.3B and 4.4B) compared to the eating response
after injection of vehicle (figure 4.3A, 4.4A). Figure 4.3B shows that three
days of DE significantly reduced the ability of NPY to elicit food intake. Two-
way repeated measure ANOVA revealed a main effect of DE (F {df 1,14}=
72.39, p < 0.001) and an interaction effect between NPY and DE (F {df 1,14}
= 95.03, p<0.001). Although feeding appeared to be partially rescued in DE
animals when the dose of NPY was increased to 1.0 pg, as both a main
effect of DE and an interaction effect between DE and NPY were significant
(F {df 1,14}= 27.14; 23.6 respectively, p< 0.001 for both, Fig 3.4A). Finally, all
DE animals ate similar amounts of food when their drinking water was
returned, whether they had been given either dose of NPY or vehicle (figure
4.3C, 4.4C).
Experiment 2-injection of NPY to the LHApf after 3 days of DE
After injection of either 0.5 or 1.0 pg of NPY to the LHApf animals ate
significant amounts of food (p<0.001, fig 4.5A, 4.6A) compared to the eating
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pre-DE food intake, demonstrate a blunted feeding response to NPY. At this
point, DE-anorexia renders the PVH almost entirely insensitive to the effects
of exogenous NPY. Even a high dose of PVH targeted-NPY injection does
not stimulate significant amounts of food intake. Because NPY to the PVH or
the third ventricle normally stimulates significant amounts of food intake
(Clark, Kalra, Crowley, and Kalra, 1984; Stanley etal., 1984; Stanley et al.,
1985) our data strongly indicates DE in some way desensitizes the PVH to
the feeding effects of NPY. Similar data have been shown in other models of
anorexia. Anorexic tumor-bearing rats, senescent rats and rats with
obstructive cholestasis have all been shown to be resistant to the feeding
effects of NPY injections into the PVH (Blanton et al., 2001; Chance et al.,
1996; Rioux et al., 2001).
Although the PVH was not responsive to the food intake stimulatory effect of
NPY, our results revealed that exogenous NPY continues to stimulate food
intake when it is directed towards the ventromedial edge of the fornix,
posterolateral to the PVH, an area commonly referred to as the LHApf
(Blanton etal., 2001; Chance etal., 1996; Rioux et al., 2001; Stanley etal.,
1993; Swanson, 1998). In fact, a high dose (1.0 pg) of NPY targeted to this
locus rescues the NPY feeding response entirely in moderately anorexic
animals. The reasons for the anatomical difference in NPY effectiveness in
DE-anorexia remains unclear. Radio-labeled peptide YY has been shown to
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bind heavily at the LHApf and this area is the most sensitive site for NPY to
elicit food intake (Stanley et al., 1993). Thus this area may have more
receptors that remain active for longer in DE. Alternatively, it has been
suggested previously that while NPY in both the PVH and the LHA stimulate
food intake, these loci may actually be responsible for different food intake-
related functions (Stanley, 1993; Stanley etal., 1993). The PVH contains
two sets of neurons: neuroendocrine neurons that regulate endocrine
functions and pre-autonomic neurons with descending projections that
oversee both autonomic and behavioral motor events (Swanson etal., 1980)
and in fact, several endocrine and autonomic effects accompany feeding
after NPY injection into the PVH including a decrease in gastric acid
secretion (Monnikes, Tebbe, Bauer, Grote, and Arnold, 2000; Tebbe,
Mronga, Schafer, Ruter, Kobelt, and Monnikes, 2003) and sympathetic
activation of brown fat (Billington, Briggs, Harker, Grace, and Levine, 1994;
Kotz, Briggs, Grace, Levine, and Billington, 1998; Kotz, Wang, Briggs,
Levine, and Billington, 2000), increased release of corticosterone and insulin
(Abe, Saito, Ikeda, and Shimazu, 1991; Gao, Ghibaudi, and Hwa, 2004;
Hanson, and Dallman, 1995; Wahlestedt, Skagerberg, Ekman, Heilig,
Sundler, and Hakanson, 1987). These effects do not appear to accompany
injection of NPY into the LHA. Rather, the LHApf has consistently been
shown to be intimately involved with the neural control of food intake. In
addition to being the most sensitive site for the orexigenic effects of NPY,
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neurotoxic and electrical lesions of the LH and the LHApf impair eating
(Anand etal., 1951; Winn, 1995) and electrical stimulation elicits eating in
satiated rats (Takaki, Aou, Oomura, Okada, and Hori, 1992) while glutamate
injections to the LHApf also regulate feeding but not other behaviors (Khan,
Curras, Dao, Jamal, Turkowski, Goel, Gillard, Wolfsohn, and Stanley, 1999).
In this sense, if NPY singularly stimulates food intake at the LHApf, its action
may remain viable for longer than the PVH, whose feeding action is
complimentary to other functions related to eating.
While the LHApf remains viable to NPY stimulatory effects for longer than the
PVH, the mechanisms activated by DE eventually inhibit the response to
NPY at the LHApf as DE-anorexia continues to progress. After five days,
DE is well-established and presumably pathways activated to preserve fluid
balance override those responsive to energy metabolism. The mechanisms
by which DE inhibits NPY output is not definitively known but potential
mediators may be the CRH neurons located within the bed nucleus of the
stria terminalis (BNST) and a subset of neurons within the LHA itself that
produce CRH and NT (Watts, 1999; Watts, 2000; Watts etal., 1995b). At
these sites, gene expression of CRH and NT begin to increase as soon as
DE-animals begin to drink hypertonic saline but before anorexia manifests.
Further, the increase in the strength of DE-anorexia correlates with the
increase in gene expression of CRH and by 24 hours after anorexia is
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reversed by returning the animal’s regular drinking water, the CRH mRNA
levels revert to normal (Watts, 1999). Central injections of NT reduce food
intake and CRH is a well-documented anorexic agent as intraventricular
injections of CRH inhibit both deprivation-induced eating and NPY-induced
feeding, and NPY feeding can be significantly increased if CRH receptors are
blocked by an antagonist (Heinrichs, Menzaghi, Pich, Hauger, and Koob,
1993; Levine et al., 1983; Morley, 1987). NPY acts through a family of
receptors which are coupled to inhibitory G-proteins that inhibit the
accumulation of cAMP (Gicquiaux, Lecat, Gaire, Dieterlen, Mely, Takeda,
Bucher, and Galzi, 2002) . The cellular effects of CRH acts in an opposite
fashion, through the stimulation of cAMP levels (Chen, Hatalski, Brunson,
and Baram, 2001). Recently investigators have reported evidence of CRH
and NPY intracellular counter-regulation through reciprocal effects on cAMP
levels (Sheriff, Dautzenberg, Mulchahey, Pisarska, Hauger, Chance,
Balasubramaniam, and Kasckow, 2001), lending further support to the
possibility that these two neuropeptides may interact with each other to effect
behaviors such as food intake. However, the fact that the injection of
substances into the LHA that increase the amount of cAMP also elicit feeding
behaviors (Gillard, Khan, Grewal, Mouradi, Wolfsohn, and Stanley, 1998)
reminds us that the mechanisms by which food intake is stimulated is
complex and probably not regulated entirely through a simple up- or down-
regulation of cAMP.
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Unlike the anatomical and DE-related differences of our first findings, the
latency to eat after NPY injection was almost identical in all the groups of
animals. Animals given intraparenchymal NPY eat, on average, within the
first ten minutes following the start of the injection whether the animal was
DE-anorexic or EU, and regardless the site of injection, the PVH or the
LHApf. Our data are almost identical to previous reports of NPY feeding
latency (Stanley et al., 1985). It is interesting to note that although latencies
are the same between EU and DE animals, the feeding rates are very
different. EU animals eat at a high rate for the first hour and continue to eat
through the remainder of the experiment. In contrast, the group of DE
animals resistant to NPY feeding stimulatory effects eat at a much lower rate
and hardly eat anything following the first hour, similar to an eating pattern of
a more-satiated animal (Marin Bivens, Thomas, and Stanley, 1998).
Critically, DE animals can and will eat. The NPY stimulus appears to
continue to stimulate meal initiation mechanisms but the sustained feeding
effects are considerably weaker particularly when DE-anorexia is well
established. In light of data suggesting NPY acts to elicit eating behavior, at
least in part, through suppressing the satiating effect of food (Lynch, Hart,
and Babcock, 1994), it seems reasonable to suggest DE mechanisms may
function to counteract this component of NPY feeding. This idea is
suggested by studies done on eating behaviors on fluid restricted humans
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(Bolles, 1969; Engell, 1988). Here, individuals who reported a increased
intensity of thirst also consumed less food despite having hedonic ratings of
food acceptability identical to that of individuals with ad libitum fluid intake.
Finally, we find that upon drinking water again, all the DE animals show a
stereotypical bout of avid food consumption in which animals eat in one hour
more than they have spontaneously eaten in the previous 12 (Watts, 1999).
All of our DE animals eat the same amount of food after anorexia reversal,
despite differences in NPY injection sites or whether the animals had
previously received an injection of NPY or not, and in fact, whether or not the
animals had eaten following the NPY injection or not. We show here that the
compensatory eating episode following drinking water in DE animals occurs
even after the DE-anorexia has been over ridden by artificial NPY
administration into the LHApf. In this circumstance, the spontaneous
circadian-driven reduction in food intake that accompanies DE is rescued by
the injection of NPY, stimulating food intake and interestingly, the animals
then eat robustly again for the hour following the initiation of drinking water.
The fact that an injection of NPY four hours previous to the reversal of
anorexia by drinking water has no effect on the amount of feeding suggests
the exogenous NPY has no role in feeding at this point. This is not surprising
in light of data showing high extracellular levels of NPY are rapidly reduced
(Kalra et al., 1991; Stanley etal., 1993). Interestingly, eating after NPY
injection into the LHApf in the hours immediately before the return of drinking
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water also has no satiating effect on the compensatory feeding bout when
anorexia is reversed. The mechanisms by which anorexia is reversed is not
clear at the present time but they appear to disinhibit a stimulatory network
to promote eating behavior quickly, powerfully and with little regard for any
previous satiating effects of food.
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CHAPTER FIVE
OREXIGENIC EFFECTS OF CENTRAL AGRP INJECTION
ARE REVERSED BY ANOREXIA
INTRODUCTION
We have previously shown that the anorexia that develops after drinking
hypertonic saline persists in the face of at least three metabolic challenges:
prolonged energy deficit, overnight starvation, and glucoprivation. In the
absence of DE, each of these energy deficient states elicits food intake
through the release of NPY in the hypothalamus that originated from cell
bodies located in the ARH or the hindbrain (Li et al., 2004; Minami etal.,
1995; Ritter et al., 2001b; Sahu etal., 1988; Sawchenko etal., 1985).
Furthermore, injection of exogenous NPY directly towards the paraventricular
nucleus and the perifornical lateral hypothalamic area also does not rescue
feeding after DE-anorexia has become well established (Salter and Watts,
2004). Together, these data suggest that DE activates mechanisms that
specifically inhibit NPY neural circuits involved with feeding, specifically at
hypothalamic sites of NPY efferent terminations.
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A significant component of the NPY neurons in the ARH also express and
co-release Agouti Related Peptide (AgRP), the endogenous antagonist for
melanocortin 3 and 4 (MC3/4) receptors (Broberger et al., 1998b; Willard,
Bodnar, Harris, Kiefer, Nichols, Blanchard, Hoffman, Moyer, Burkhart, Weiel,
and et al., 1995). The melanocortin (MC) system has been shown to have
an important role in the regulation of food intake and body weight. The proÂ
opiomelanocortin (POMC) derived-peptide, a-melanocyte-stimulating
hormone (a-MSH) is the endogenous agonist of this system (Adan, Cone,
Burbach, and Gispen, 1994). Although POMC and AgRP/NPY producing
neurons are located in separate distinct regions within the ARH, their fiber
tracts project to many of the same areas of the brain where they jointly
participate in the regulation of feeding behaviors (Bagnol, Lu, Kaelin, Day,
Ollmann, Gantz, Akil, Barsh, and Watson, 1999; Elias et al., 1998b; Haskell-
Luevano, Chen, Li, Chang, Smith, Cameron, and Cone, 1999). a--MSH
binds to MC3/4-R and limits food intake while AgRP potently antagonizes
these actions (Tsujii, and Bray, 1989). In fact, a single third ventricular
injection of AgRP has been shown to increase food intake for as long as a
week (Hagan, Rushing, Pritchard, Schwartz, Strack, Van Der Ploeg, Woods,
and Seeley, 2000).
Because NPY and AgRP are produced in the same ARH neurons that are
sensitive to energy deficit (Broberger etal., 1999; Hahn, Breininger, Baskin,
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and Schwartz, 1998; Mizuno, Makimura, Silverstein, Roberts, Lopingco, and
Mobbs, 1999a; Mizuno, and Mobbs, 1999b) we would expect these peptides
to receive the same afferent regulation. However, AgRP activates feeding
through the MC3/4 receptor system, a system separate from that of NPY.
Therefore, examining AgRP gene expression and AgRP related-feeding
behavior in DE will provide information on how adaptive mechanisms might
function in anorexia.
MATERIALS AND METHODS
ANIMALS AND PROCEDURES
Adult male Sprague-Dawley rats (240-260 g) obtained from Harlan
laboratories were individually housed in suspended Plexiglass cages with
sanitized wood chips. They were maintained in a temperature-controlled
room on a 12:12-h light-dark schedule with lights on at 0600. Rats were
provided continuous access to food (Teklad rodent diet 8604) and water
throughout the experiment, except where stated. In some animals, drinking
water was replaced with during 2.5% saline solution for three days. Body
weights and food intake were measured through out the experiment. All
procedures have been approved by the local institutional animal care and
use committee.
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SURGICAL PROCEDURES
Rats were handled daily for approximately 5 days before any surgical
intervention and daily thereafter. Guide cannulas were fabricated from 26
gauge stainless steel tubing and occluded with a removable stainless steel
obturator. For stereotaxic implantation the guide cannula, rats were
anesthesized with an injection (100pil/kg i.m.) of ketamine and
xylazine/acepromazine cocktail solution (5 ml ketamine HCI.100 mg/ml; 2.5
ml xylazine, 20mg/ml; 1 ml acepromazine, 10 mg/ml; 1.5 ml 0.9% saline
solution). The skull was exposed and trephined at the implantation site using
flat-skull coordinates of 2.1 mm caudal to Bregma at midline and 6.8 mm
ventral to the dura for injections into the third ventricle. Guide cannula were
lowered to the desired site and anchored to the skull with dental acrylic and
stainless steel screws. After the skin incision was sutured, rats were given an
injection of analgesic (Banamine HCI, 2.5 mg/ml, 10Opil/kg i.m.), allowed to
recover from anesthesia and were then returned to a clean home cage.
Behavioral testing began 7-9 days post-surgery, when body weight and food
intake had returned to pre-surgical values.
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INJECTIONS
All injections were given between 0900-1000 h in equal volumetric doses of 2
pi over 60-120 seconds. The obturator was removed and replaced with a 33
gauge stainless steel injection cannula attached by polyethylene tubing to a
25 pi Hamilton syringe. During injections, the movement of a small bubble in
the calibrated infusion line was used to verify injectate delivery. In addition,
the system was tested before and after each injection to ensure it was free of
leaks or plugs.
FEEDING AND FLUID INTAKE TESTS
After each injection, the obturator was replaced and animals were returned to
their home cages with water or 2.5% saline to drink and a pre-measured
amount of rat chow. Food consumption was measured to the nearest 0.1 g
by weighing food and crumbs remaining in cages each hour for the amount
specified. Feeding latency was also measured. A pre-measured amount of
food was placed in the test cage each day and food consumption was
measured by weighing the food remaining in the cage to the nearest 0.1 g
each. Similarly, fluid intake was measured by placing a pre-weighed water
bottle containing 2.5% saline or water in the test cage. At the conclusion of
the time period being investigated, the bottle was re-weighed and the
decrease in bottle weight was considered milliliters drank by the animal.
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Nocturnal and diurnal food and fluid intake were measured twice daily
between 07.00-08.00 and 16.00-17.OOh for three days following injections.
EXPERIMENTAL TIMELINE
Prior to any AgRP injection, cannula placement into the 3rd ventricle was
verified by a drinking test to 10 ng angiotensin II in vehicle while animals
were water replete. Only rats that drank a minimum of 5 ml of water within
the hour following angiotensin II injection were included in further
experimentation. Prior to AgRP injections, nocturnal and diurnal food and
fluid intakes were measured for three days to obtain each animals baseline
consumption. Rats were then weight-matched and placed into groups. For
DE groups, drinking water was replaced with 2.5% saline whereas EU
weight-matched control animals remained on drinking water throughout the
experiment. 24 hours after their drinking water was replaced with 2.5%
saline, DE rats were injected with one of 3 different doses (0.1, 0.5, or 1.0
nMol) of the amidated COOH-terminal AgRP fragment (83-132)-NH2
(Quillan, Sadee, Wei, Jimenez, Ji, and Chang, 1998)( Phoenix
Pharmaceuticals) or vehicle. Each EU rat was injected in the same manner
directly following that of its weight-matched (pre-dehydration) DE partner.
Food and fluid intake was monitored for three hours following injection and
for the next three days. On the morning of the 4th day, DE animals' drinking
water was replaced and for the following hour, food and fluid intake was
117
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tested. A separate brief experiment was done on 5 EU-AgRP injected
animals(1.0 nMol dose) to investigate if AgRP stimulated salt-appetite.
These animals were given two water bottles, one with a 1.9% saline solution
and one with regular drinking water and the bottle placements were rotated
every two days. Fluid intakes were measured twice daily, between 07.00-
08.00 and 16.00-17.O O h for three days following injections.
! , | diurnal and
Food and Fluid | 1h , 1-3h i nocturnal intakes i ^
intake meaured 1 I 1 1 ! 1 1 1
1 --------------------------------------------------------------------------1 -------------------- 1 ----------------------------------------- 1 —
| 7-9 days j 2 days j 24 hours | 3 days
Cannulate angiotensin Dehydration ^W at e r returned
drinking (DE groups) AgRP or Vehicle t0 g r o u p s
test or ad lib water
(EU groups)
Figure 5.1: Timeline for fluid and food intake experiments for animals given a
3V injection of vehicle or one of three doses of AgRP (0.1, 0.5, or 1.0 nMol).
Animals were given the first injection of VEFI or AgRP 7-9 days following
surgery. Animals were placed into groups, two groups being DE and one
group continuing on ad lib water intake. Each EU animal then received an
injection of AgRP, and DE animals an injection of either AgRP or vehicle the
following day. At the end of the DE period (3) all animals were tested for
their feeding response to either AgRP or VEFI and EU animals to AgRP. DE
animals were given water back at the conclusion of the four hour test.
IN SITU HYBRIDIZATION
Drinking water was replaced with 2.5% saline for 5 days in a group of rats
and food intake was measured twice daily between 07.00-08.00 and 16.DO -
17.OOh. Another group of animals was maintained on drinking water. These
118
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animals were weight-matched to the animals in the DE group. They were
then given an amount of food at the beginning of each light and dark period
equal to that eaten by DE animals (Watts et al., 1999), see chapter two).
The amount of food was calculated for each rat as a percentage of the food
eaten per 100 g of mean body weight (for two days before the beginning of
the food restriction). At the end of the five day period, rats were euthanized
with halothane anesthesia and rapidly decapitated. Brains were removed
from the skull and immersion-fixed for three days on ice cold potassium
buffered saline/4% paraformaldehyde, frozen on dry-ice and stored at -70°C.
8 series of 1-in-12 16 urn thick frontal sections were cut through the
hypothalamus. A single section series was mounted and thionin stained
while serial sections were mounted and 2 series were processed for
subsequent hybridization using S3 5 -labeled cRNA probes for AgRP (xx) and
NPY a 287 bp fragment encoding part of exon 2, (Larhammar et al., 1993),
mRNAs synthesized as previously described in Watts and Sanchez-Watts,
1995. Briefly, sections were prehybridized, hybridized for 20-22 hr at 60°C
using a probe concentration of 5x106 cpm/ml of hybridization buffer,
followed by posthybridization with RNase treatment at 37°C and room
temperature washes of 4x through 0.1x SSC, then dehydrated in alcohols.
After hybridization, sections were exposed to Microvision-C x-ray film
(Sterling Diagnostic Imaging, Newark DE) for varying periods (22 h to 3
119
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days), dipped in nuclear track emulsion (Kodak NTB-2, diluted 1:1 with
distilled water) and exposed for 2 days, developed and counterstained with
thionin. Anatomically defined regions on the film image were identified by
careful reference to local cytoarchitectonics on adjacent thionin-stained
sections and corresponding dipped autoradiographs. Photographs were
taken of corresponding Microvision C x-ray films using a SC501 CCD
camera.
STATISTICAL ANALYSIS
Data are expressed as means ± SEM. Food consumption, fluid intake and
weight changes AgRP studies were compared using one way ANOVA.
Bonferroni's pairwise multiple comparisons test was used to measure
individual differences, with critical level for significance set at p < 0.05.
RESULTS
NPY and AgrP mRNA
NPY mRNA and AgRP mRNA in pair-food restricted and DE animals is
shown in Fig 5.2. NPY mRNA expression in the ARFI was increased after 5
days of DE and was virtually identical to the increase seen in EU pair-fed
food-restricted animals. This increase was not seen in EU animals fed ad lib.
AgRP mRNA expression followed the same pattern as that seen with NPY; a
120
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noticeable increase in both DE-anorexic and EU-FR over the mRNA
expression seen in EU ad lib fed animals.
Effects of AgRP on food intake in DE and EU animals
While the lowest dose of AgRP in EU animals did not significantly increase
food intake 3 hours after injection over the amount of food eaten by vehicle
injected animals, both the higher doses (0.5 and 1.0 nMof) elicited significant
amounts of food intake (Fig 5.3, F{df 7,59}=10.2, p< 0.001). None of the
doses of AgRP increased food intake in DE animals over the amount of food
eaten by both DE and EU animals injected with vehicle (Fig 5.3).
A single AgRP injection to the 3v of EU animals elicited a dose-responsive
increase in 24 hour food intake over that seen in EU vehicle-injected animals
(Fig 5.4). The 0.1 nMol dose of AgRP elicited a significant increase in food
intake only in the first 24 hours after injection (F {df 3,24}= 20.80, p<0.001)
whereas both 0.5 and 1.0 nMol doses of AgRP significantly increased food
intake in each of the 3-24 hour periods measured (p<0.001 for each). None
of the doses of AgRP increased intake in DE animals over that seen in
vehicle-injected DE-anorexic animals. In fact, the higher doses significantly
intensified the DE-anorexia in the last two 24 hour increments of food
monitoring (p< 0.001).
121
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Effects of AgRP on body weight in DE and EU animals
A single 1.0 nMol dose of AgRP significantly increased the amount of body
weight EU rats gained over that by EU rats injected with vehicle
(F{df3,24}=9.66, p<0.001, Figure 5.5). The body weight increase seen after
the 0.5 nMol dose of AgRP approached but did not reach significance
whereas the lowest dose did not change the amount of body weight gain in
control animals.
Conversely, AgRP significantly exacerbated the amount of body weight loss
that normally accompanies DE (F{df 3,28}=6.78.54, p<0.01, Figure 5.5).
Specifically, both the 0.5 and 1.0 nMol doses caused accelerated weight loss
in DE rats.
Effects of AgRP on fluid ingestion in DE and EU animals
When fluid was measured for the three hours immediately following injection,
the 1.0 nMol dose of AgRP elicited significant amounts of water intake (Fig
5.6, F{df 3,25}=11.54, p<0.001) over the amount vehicle-injected rats drank,
but none of the doses of AgRP increased fluid intake significantly in DE rats.
However, in DE rats a 0.5 or 1.0 nMol dose of AgRP had a significant effect
122
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EU ad lib
A B
c D
E
3v
F
j \ ~ • '
- f c ^ r
EU- Food restricted
G
* .
H
i J
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1
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DE
M N
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Figure 5.2: NPY and AgRP mRNA
hybridizaton in the arcuate nucleus
of the hypothalamus.
Bright field photomicrographs of
NPY (A.B.G.H.M.N) and AgRP
(C,D,l,J,0,P) mRNA hybridization
in the ARH at approximately levels
28 (AIC,E,G,l,KJ M ,0,Q )or31
(B.D.F.H.J.L.N.P.R) of Swanson
Rat Brain Atlas (1992). Shown are
serial sections from a control
animal allowed ad lib food
availability (A-F), an euhydrated
food-restricted animal (G -L ), or an
animal after 5 days of dehydration
(M-N).
E,F,K,L>Q,R show serial sections
stained with thionin for anatomical
reference.
3v, third ventricle.
123
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0
VEH 0.1 0.5 1.0
Dose AgRP (nMol)
Figure 5.3: Hypertonic saline ingestion inhibits feeding 3 hours after 3v
AgRP injections.
Mean cumulative intake( ±SEM) 3 hours after injection of different doses of
AgRP in EU animals (open circles) or animals given 2.5% saline for 24
hours prior to injection( closed circles in gray background). See text for
statistical significance.
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1 6 0
140
CD
Euhydrated
animals
.9 2 . 120
100
TJ
80
60
Dehydrated
animals
40
20
prior 24 Day 1 Night 1 Day 2 Night 2 Day 3 Night 3
Figure 5.4: DE inhibits feeding in response to 3v AgRP injection.
Mean nocturnal and diurnal cumulative intake( ±SEM) after injection of
different doses of AgRP in EU ( vehicle, open circles; 0.1 nMol, open
squares; 0.5 nMol, open triangles and 1.0 nMol, open diamonds) and DE
animals (vehicle, closed circles; 0.1 nMol, closed squares; 0.5 nMol, closed
triangles and 1.0 nMol, closed diamonds). See text for statistical
significance.
125
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35.0
25.0
15.0
3
c
Id 5.0
C O
c o
n
a >
o >
c
co
s z
O
-5.0
-15.0
-25.0
-35.0
-45.0
Euhydrated
animals
Dehydrated
animals
0 24 48
Hours after Injection
72
Figure 5.5: Dose-responsive changes in body weight after 3v injection of
AgRP are different in EU and DE animals.
Change in body weight from baseline ( ±SEM) over 72 hours following a
single injection of vehicle or one of three different doses of AgRP in EU
(vehicle, open circles; 0.1 nMol, open squares; 0.5 nMol, open triangles and
1.0 nMol, open diamonds) and DE animals (vehicle, closed circles; 0.1 nMol,
closed squares; 0.5 nMol, closed triangles and 1.0 nMol, closed diamonds).
See text for statistical significance.
126
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on drinking of hypertonic saline over three days (Figure 5.7, F{df
3,26}=14.76, p<0.001). EU rats injected with either 0.5 or 1.0 nMol AgRP,
but not 0.1 nMol, also had a significant increase in 72 hour intake of water
over that of vehicle-injected EU (p<0.001 for both). EU AgRP-injected rats
(1.0 nMol) given a two- bottle preference test clearly preferred regular
drinking water to a mildy hypertonic saline solution, drinking 122.3 ± 0.8 mis
of water over the three days of testing vs. 5.2 ± 0.2 mis of saline.
Effects of AgRP on compensatory feeding after water returned to DE animals
Three days after AgRP was administered to DE rats the hypertonic saline
was replace with drinking water and all animals demonstrated a
compensatory bout of feeding within one hour (Figure 5.8). However, the
feeding response of DE animals that had previously received an injection of
either 0.5 or 1.0 nMol AgRP, but not 0.1 nMol, ate significantly less than
vehicle injected control animals (p<0.001). The amount of water the animals
consumed during this test was not different between groups as 0.1, 0.5 or 1.0
nMol AgRP-injected animals consumed 25.2 ± 2.5, 27.5 ±2.3, and 28.3 ±1.3
mis of water, respectively. Vehicle-injected DE rats consumed 26.7± 1.6 mis
of water in the one hour after water was returned.
127
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12
10
8
6
2
0
VEH 0.1 0.5 1.0
Dose AgRP (nMol)
Figure 5.6: A high dose of AgRP stimulates fluid intake in EU, but not DE
animals, 3 hours following a single 3v injection.
Mean cumulative fluid ingestion( ±SEM) 3 hours after injection of different
doses of AgRP in EU animals drinking water (open circles) or DE animals
drinking 2.5% saline (closed circles). DE-animals were given hypertonic
saline to drink for 24 hours prior to injection. See text for statistical
significance.
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Veh 0.1 0.5 1.0 Veh 0.1 0.5 1.0
Dose of AgRP (nMol)
Figure 5.7: 72 hour fluid intake after different dose AgRP injection in EU and
DE rats.
A) Mean (±SEM) total cumulative water intake in EU rats 72 hours after
vehicle or one of three doses of AgRP. B) Total cumulative hypertonic
saline intake 72 hours after vehicle of AgRP injection in DE rats. DE rats’
water was replaced with 2.5% saline 24 hours prior to the injection. See text
for statistical significance.
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aCSF 0.1 nM 0.5 nM 1.0 nM
AgRP AgRP AgRP
Figure 5.8: Food intake 1 hour after water is returned to DE animals.
Mean (±SEM) food intake in DE animals one hour after drinking water was
returned. Water was returned three days after injection of vehicle or one of
three doses of AgRP. See text for statistical significance.
DISCUSSION
Our present results demonstrate four main points in regards to how AgRP
functions during DE-anorexia. First, both AgRP and NPY mRNA are up-
regulated in DE-anorexic and pair-fed animals compared to EU control
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animals that are allowed to eat ad-lib. Second, the anorexia and weight loss
that accompanies DE is not reversed by a central injection of AgRP. Rather,
at 48 hours post-injection, the anorexia is more pronounced than that seen in
control animals. Third, AgRP injections markedly stimulate saline ingestion
in DE-anorexic animals, which in turn likely exacerbates their anorexia.
Finally, all DE animals eat when drinking water is returned, regardless of
receiving a prior vehicle or AgRP. However, animals receiving an AgRP
injection 72 hours earlier eat significantly less than those animals that had
received a vehicle injection, despite drinking similar amounts of water.
Both DE-anorexic and hungry animals demonstrate profiles of
diminished energy status including reduced plasma leptin and insulin, and
elevated corticosterone, and these signals most likely initiate the increased
ARH NPY mRNA seen after both these metabolic challenges (Watts, 1999;
Watts et a l, 1999). Here we show that AgRP mRNA is increased in a
manner virtually identical to the increase seen with NPY expression.
Because AgRP is co-expressed with NPY in a population of ARH neurons
expressing leptin receptors and demonstrating sensitivity to energy status
(Hahn etal., 1998; Korner, Savontaus, Chua, Leibel, and Wardlaw, 2001;
Mizuno etal., 1999a; Mizuno etal., 1999b) the same afferent inputs are likely
responsible for the increase in both sets of mRNA. Both these
neuropeptides share some of the same efferent targets, particularly the PVH
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and LHA, (Broberger etal., 1998b; ELIAS etal., 1998b) and both stimulate
food intake (Rossi, Kim, Morgan, Small, Edwards, Sunter, Abusnana,
Goldstone, Russell, Stanley, Smith, Yagaloff, Ghatei, and Bloom, 1998;
Stanley, Magdalin, Seirafi, Nguyen, and Leibowitz, 1992). However, they do
so with different receptor systems. NPY utilizes a family of inhibitory G-
protein-coupled receptors whereas AgRP functions largely to block agonists
from accessing MC3/4 receptors. Therefore, because DE-anorexic and food-
restricted hungry rats demonstrate nearly identical hypothalamic mRNA
expression but demonstrate divergent feeding behaviors, it is possible that
the efferent targets of AgRP and NPY projections respond differently in DE.
In support of this idea, we have previously shown exogenous NPY injections
into terminal sites that stimulate food intake in EU rats will no longer be as
effective in doing so when rats are DE (see chapter four). We now show that
exogenous AgRP also does not stimulate food intake in DE rats.
Interestingly, this occurs even when the AgRP is injected as few as 24 hours
after rats were started on hypertonic saline, a time when plasma osmolality is
not significantly elevated beyond normal values (Watts et al., 1995a).
Furthermore, despite that a single injection of AgRP induces a long-term
(3d) increase in food intake and weight gain in EU, it does not stimulate food
intake in DE animals. In fact, the anorexia and weight loss that normally
accompanies DE is intensified by AgRP.
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The mechanisms that lead to an increased intensity of anorexia and weight
loss after exogenous AgRP in DE animals is unclear. Elevations of leptin or
insulin in the brain are associated with decreased food intake (Friedman et
al., 1998; Woods etal., 1979). Central AgRP injections have been shown to
increase plasma leptin and insulin concentrations independent of
hyperphagia and fat-pad weights (Korner, Wissig, Kim, Conwell, and
Wardlaw, 2003; Small, Kim, Stanley, Mitchell, Murphy, Morgan, Ghatei, and
Bloom, 2001). DE-anorexic rats have undetectable levels of leptin or insulin
(Watts et al., 1999) thus even small elevations of either of these hormones
could increase the intensity of anorexia in these animals. Alternatively, the
fact that DE animals given AgRP increase considerably their hypertonic
saline consumption likely exacerbates anorexia, as food intake decreases
linearly as drinking saline progresses (Watts, 1999). The reasons for the
dramatic increase in saline consumption in these AgRP-injected DE rats is
unclear, however it is not due to a stimulation of salt appetite as AgrP-
injected EU animals display a clear preference for drinking water over a
mildly hypertonic saline solution. EU rats did increase their fluid intake after
AgRP injection over that consumed by vehicle-injected control animals.
Thus, it is possible AgRP has dipsogenic effects that are markedly
unmasked in the face of the hydrational challenge poised by DE. Rather
than excusively targeting forbrain mechanisms, it is important to consider the
notion caudal hindbrain (HB) substrates may mediate at least some of the
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behavioral alterations seen after AgRP injection in DE animals. The dorsal
motor nucleus of the vagus nerve and the adjacent commissural nucleus of
the solitary tract (cNTS) house the highest MC4 receptor density found in
the brain (Mountjoy, Mortrud, Low, Simerly, and Cone, 1994) and injection of
MC-R ligands directly into the HB affect feeding and weight changes in a
manner qualitatively similar to that seen after injections into the forebrain
(Grill, Ginsberg, Seeley, and Kaplan, 1998). Furthermore it is possible that at
least some of our third ventricular AgRP injections traveled to the HB through
the normal directional flow of cerebral spinal fluid. Two other lines of
evidence support the notion that HB substrates may participate in the
behavioral alterations seen after AgRP application during DE. First,
however, it is important to recognize that although POMC neuronal
populations are found in both the forebrain and the hindbrain (Bronstein,
Schafer, Watson, and Akil, 1992; Palkovits, Mezey, and Eskay, 1987), AgRP
has thus far only been found to be expressed in the ARH (Broberger et al.,
1998b). Furthermore, there are few documented AgRP projections to the HB
(Haskell-Luevano et al., 1999) so it is possible that the presence of large
amounts of exogenous AgRP in the hindbrain is not physiologically relevant.
Rather, in the face of physiological challenge presented by DE, the agonist
for MC3/4 receptors, a-MSH, may be a predominate factor underlying the
normally decreased food intake in this circumstance. There is a large
population of POMC neurons that produce the anorexic a-MSH located
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within the cNTS, there is dense POMC innervation to the MC4 receptors in
the HB (Bronstein et al., 1992; Palkovits et al., 1987) and application of MC
receptor agonists to this area elicits a reduction of food intake (Grill et al.,
1998). Second, the area postrema is an important mediator in
osmoregulation and NaCI intake (Strieker, Craver, Curtis, Peacock-Kinzig,
Sved, and Smith, 2001) and has bidirectional connections with the cNTS.
Therefore substrates within the HB are positioned to participate in some way
to osmoregulation and the subsequent metering of osmole intake during DE,
in either the form of food or NaCI. Therefore it is possible that the abnormal
introduction of high levels of exogenous AgRP into this system during DE
may disrupt in some way the normal homeostatic mechanisms that operate
in conjunction with forebrain osmoreceptors to limit the addition of salt loads
in already DE animals.
This idea of HB involvement is particularly interesting in light of our data
showing the anorexia manifested by DE-rats is not fully reversed by drinking
water in rats that had received a prior AgRP injection, despite that these
animals drink similar amounts as vehicle injected control animals. Evidence
suggests the resolution of DE-anorexia after drinking water occurs too
rapidly to be the result of an amelioration of plasma hyperosmolality. Rather
osmoreceptors located in the lower stomach and upper duodenum that signal
through the vagus nerve may be responsible (Schoorlemmer, and Evered,
135
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2002; Strieker, Callahan, Huang, and Sved, 2002; Strieker, Huang, and
Sved, 2002). Therefore in light of data showing AgRP mediates long-term
(up to 7 days) neuronal alterations (Hagan etal., 2000), it is tantalizing to
speculate that during DE, high doses of AgRP alters the animal’s ability for
appropriate osmoregulation and that this deficiency is not reversed by
drinking water due to the long term effects of AgRP acting on substrates
within the HB.
Possibly, the curious results presented in this chapter present more
questions than they have answered in regards to the organization and
interactions of neural circuits involved in adaptive mechanisms in DE. Does
AgRP serve as a primary dipsogen? Does AgRP intensify anorexia through
receptor mediated mechanisms and an increased circulating levels of leptin
and possibly insulin? Rather, is the increased anorexia specifically a
consequence of increased hypertonic saline consumption? Can the results
presented here be duplicated if AgRp injections are constrained to the 4th
ventricle? These questions present several lines of investigation
opportunities to be carried out in our laboratory in the future.
Regardless, several recent investigations have suggested AgRP to be an
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
agent that simply increases food intake. However, the results we
demonstrate here illustrate that mechanisms underlying adaptive feeding
behaviors in energy homeostasis are rarely simple or straightforward.
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CHAPTER SIX
CONCLUSIONS
Stimulatory circuits
In most mammals the levels of body energy stores remains relatively
constant for long periods of time. This occurs despite that daily energy
intake and expenditure are relatively stable from day to day. If energy intake
is decreased and body weight is lost, systems are set into motion for
increased food intake and body weight stores are returned to normal (Bray et
al., 1998; Keesey et al., 1997). This is the case, even if the body weight lost
is in the form surgical excision of fat stores. Strikingly, the individual who has
lost weight to the scalpel will eat sufficient extra calories to gain enough
weight to return to the previous pre-surgical weight (Faust, Johnson, and
Hirsch, 1977a; Faust, Johnson, and Hirsch, 1977b). Indeed, whether energy
deficit is prolonged with extensive weight loss or as acute as a missed meal,
the regulatory system responsible for maintaining energy homeostasis is
reliably quick and powerful. Peripheral signals of energy status are
transduced into appropriate motor patterns to favor energy consumption.
Mounting evidence suggests that neuronal pathways in the hypothalamus
138
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are key substrates in the refeeding after energy deficit, primarily those
pathways that utilize NPY in the ARH-PVH-LHA circuit (Figure 6.1).
Inhibitory circuits:
Anorexia reduces or eliminates the behavioral drive to increase energy intake
in the face of negative energy balance. The anorexia that accompanies DE
demonstrates numerous hallmark signs of energy deficit; body weight loss,
diminished circulating leptin and insulin, and increased circulating blood
glucocorticoids, in addition to increased NPY gene expression in the ARH
((Watts et al., 1999). We have presented the results of a series of
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Y/NE/E)
FEEDING
Figure 6.1: Circuit diagram illustrating neural substrates involved in
refeeding after a period of negative energy balance. Signals of decreased
energy status; hypoglycemia, decrease in circulating leptin, decrease in
circulating insulin, or increased circulating glucocorticoids all stimulate
increased production and release of NPY in the hypothalamus. Through
complex and distributed neural circuits, these neurons initiate mechanisms
that increase drive to eat.
139
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experiments designed to investigate how the mechanisms that normally elicit
feeding in response to negative energy balance can be inhibited.
Cumulatively, our data strongly suggest the inhibition that accompanies DE
targets the output of those insulin and leptin-sensitive NPY neural
mechanisms that initiate behavioral motor actions to increase food intake.
By using the competitive glucose analogue 2DG as a tool to delineate the
neural circuits underlying anorexia, we show the inhibition that accompanies
DE is specific to those stimulatory pathways that activate feeding. The
complimentary glucoregulatory responses that manifest after 2DG
administration are maintained while feeding in DE is not, suggesting those
feeding pathways utilizing NPY are inhibited.
Furthermore, neuronal activation after 2DG administration in DE suggest
those areas that receive afferent information regarding energy status
continue to respond in DE. However, those areas where NPY stimulates
food intake show a decreased activation after 2DG in DE. Finally, we show
that DE progressively inhibits the sensitivity to NPY activity, as animals
demonstrate reduced responses to both overnight starvation and exogenous
intracerebral NPY administration over time during DE. The fact that NPY
insensitivity is shown specifically at the PVH and LHA, areas that receive
NPY afferents arising from two separate neural circuits sensitive to energy
status, further supports the notion the afferent activity is intact in this model.
140
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Thus our data supports the hypothesis that the anorexia that manifests with
DE results, at least partially, from an inhibition in the post-synpatic cellular
cascade rather than as a result of inhibited release of NPY (Figure 6.2).
In fact, DE-anorexia appears to be generated by activity in separate inhibitory
circuits. These circuits may involve neurons that produce anorexigenic
neurpeptides such as CRH, neurotensin or oxytocin found in those parts of
the LHApf, PVH, and bed nucleus of the stria terminalis that are targeted by
Osmolality
Leptin
Glucocorticoid
i ypocflycerrua
â–º LHA
NPY/AaRP I
► ARH ►• PVH
â–º CORTEX
* PAG
- -â–ºHINDBRAIN
CATECHOLAMINE
NEURONS
â–º PBN
Potential sites of interaction for generating DE-anorexia
Anorexia
Figure 6.2: Circuit diagram illustrating neural substrates involved in
refeeding after a period of negative energy balance continue to stimulate
increased production and release of NPY in the hypothalamus. Separate
circuits activated by increased plasma osmolarity signal through the LHA to
inhibit the post-synaptic activity of NPY efferent information, namely at the
PVH and LHA, resulting in anorexia.
141
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plasma osmolality-sensitive rostral hypothalamic afferents (Heinrichs etal.,
1993; Kelly et al., 1996; Levine etal., 1983; Olson, Drutarosky, Strieker, and
Verbalis, 1991a; Olson, Drutarosky, Strieker, and Verbalis, 1991b; Watts et
al., 1999).
Disinhibitory circuits:
The DE-anorexia that develops progressively over 3-5 days of drinking
hypertonic saline is reversed quickly and robustly upon the return of drinking
water. The manner in which these DE-animals drink and eat during this time
is temporally ordered, so the rat almost always drinks for 7-10 minutes and
then switches to eating voraciously (Watts, 1999). Presumably the main
priority is first to restore the fluid compartment and then to replenish energy
stores. With the exception of those rats injected with AgRP (see chapter
five), all the animals administered orexigenic agents to stimulate eating prior
to the restoration of drinking water ate approximately the same amount of
food as control DE animals. This may be simply the result of upper
limitations of gastric distensibility as animals drink significant amounts of fluid
followed by the consumption of more food in one hour than the rats had
consumed in the previous 12 (Watts, 1999). Alternatively, the highly
stereotypical compensatory feeding bout may be driven by a separate circuit
in which orexigenic hypothalamic neuropeptides do not have a prominent
role, at least not immediately after initial water drinking. Because the
142
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behavioral switch from drinking to eating occurs before plasma
hyperosmolality is normalized, the sensors responsible for initiating this
circuit is may, in fact, be located within the gut. Slow infusions of water into
the stomach and duodenum, but not the portal vein, of fluid-challenged rats
not allowed to drink was reported to reverse DE-anorexia (Schoorlemmer et
al., 2002). The fact that arginine vasopressin (AVP) secretion in dehydrated
dogs (Appelgren, Thrasher, Keil, and Ramsay, 1991) and rats (Huang, Sved,
and Strieker, 2000) is inhibited by drinking water before significant reductions
in plasma hyperosmolality supports this notion that centrally regulated motor
events can be modified by drinking hypotonic solutions. Further,
oropharyngeal mechanisms may also be involved because AVP is reported
to quickly (< 5 min) decrease when dehydrated humans (Figaro, and Mack,
1997) and dogs (Thrasher, Nistal-Herrera, Keil, and Ramsay, 1981) drink but
the swallowed fluid is immediately removed by tube or fistula. Thus, we
hypothesize that the quick and enthusiastic reversal of DE-anorexia arises
from disinhibitory circuits initiated by sensors gut and possibly oropharyngeal
sensors (Figure 6.3).
Although investigating the neural mechanisms that underlie DE-anorexia will
not reveal all of the various mechanisms responsible for different types of
anorexia, it does provide a useful paradigm with which approach two
significant problems. First, this work should inform how ingestive behaviors
143
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are controlled at a neural systems level. The fact that there are distinct
circuits that stimulate, inhibit and switch feeding behaviors provide a unique
Osmolality â–º LHA
H y p o () I y cs m 1 8 - - -
jcoids
► n iiN U D r v M iY —
NEURONS
(AP/DMX/NTS)
CORTEX
Rcfccding
Gut Osmoreceptors
Oropharyngeal mechanis
Figure 6.3: Circuit diagram illustrating neural substrates involved in
refeeding after a period of negative energy balance. Separate circuits
activated by drinking water signal through the caudal hindbrain to disinhibit
those pathways stimulatory for feeding, resulting in a compensatory episode
of food intake.
opportunity to probe the organization of the neural networks that control
feeding. Second, DE-anorexia is particularly useful to study those circuits
that inhibit normal feeding responses to starvation. In this way, we can
make significant progress towards understanding, at a systems level ,the
neural basis of energy intake and homeostasis.
144
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Neural substrates of anorexia
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