LEPTIN RESISTANCE REDUCES GROWTH HORMONE SECRETION AND
CONTRIBUTES TO THE PATHOGENESIS OF OBESITY
By
ROBIN LEIGH MARTIN
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2000
For Jarod
ACKNOWLEDGEMENTS
There are many people to whom I wish to express gratitude for standing by me
during my efforts to get my Ph.D. First, my family: husband, George Martin; step-son,
Joshua Martin; mother, Nikki Picking; father, Tom Picking; brother and sister-in-law,
Reed and Kelly Picking, and grandparents, Don and Maxine Picking. They have all been
tremendously understanding and supportive, and I greatly appreciate it. I would also like
to acknowledge the support I have received from in-laws, aunts, uncles, cousins, nieces,
and friends, all of whom have taken the good with the bad.
I want to thank my mentor, Dr. William Millard, for his guidance and patience in
helping me reach my goal and for his faith in my "independence." Sometimes he had
more faith than I did. I would also like to thank the other members of my supervisory
committee, Dr. Ralph Dawson, Dr. Maureen Keller-Wood, Dr. James Simpkins, and Dr.
Pushpa Kalra, for their valuable advice and for allowing liberal laboratory access and use
of equipment. In addition, I thank Dr. Jeff Hughes, Dr. James Simpkins, and Dr.
Maureen Keller-Wood for finding extra work for me to do, such as performing surgeries
and running assays, when my finances were low. I am sure they benefitted from my help,
but not nearly as much as I benefitted from their money.
I wish to thank Evelyn Perez, Feng Li, DeeAnn Dugan, and Amanda Crews,
students who worked in my laboratory, for patiently allowing me to hone and sharpen my
training and supervisory skills. It was fun. Thanks also goes to Dr. Yun-Ju He and
Eileen Monck, who were quick to share their extensive technical expertise with me. I am
iii
grateful to the office staff, Donna Walko, Milena Palenzuela, Gwen Daniels, and
especially Theresa Jones, for all of their help in areas about which I know little.
I also appreciate the friendships I made with other students with whom I went
through graduate school both in my department and from other departments: Dr. Darren
Roesch, Dr. Kelly Gridley, Dr. Ming Hu, Dr. Pini Orbach, Dr. Scott Purinton, Tony
Smith, and Amanda Crews. I want to give special mention to Dr. Pattie Green, who was
a great study partner and friend, especially during our first two years as graduate students.
In addition I want to thank Dr. Bruce Jung for all the challenging, and often therapeutic,
racquetball games. The most important person I must mention, however, is Dr. Baerbel
Eppler. I am thoroughly indebted to BB for her scientific help as well as for her
friendship.
I would like to thank my friends Tim and Anita Harvey, who very unselfishly
provided me with a place to live and who took care of me when I first arrived in
Gainesville. I am also grateful to have secured friendships with Dr. Joanna Peris, Dr.
LeighAnn Stubley, Dr. Baerbel Eppler, and Theresa Jones while living in Gainesville.
Finally, I especially thank all my family and friends in Clearwater, who are awaiting my
return.
iv
TABLE OF CONTENTS
Chapter
Page
ACKNOWLEDGEMENTS
iii
LIST OF FIGURES
ix
LIST OF TABLES
xi
1 LITERATURE REVIEW
1
Obesity ........................................................................................................................ 1
Obesity Models ........................................................................................................... 3
Theories of Food Intake .............................................................................................. 6
Leptin .......................................................................................................................... 9
Roles of Leptin.......................................................................................................... 12
Regulation of Leptin ................................................................................................. 15
Leptin and Hormonal Interactions ............................................................................ 16
Leptin Receptors ....................................................................................................... 19
Leptin Signaling........................................................................................................ 23
Leptin Resistance ...................................................................................................... 25
Human Leptin Mutations .......................................................................................... 28
Leptin Treatment in Humans and Leptin Gene Therapy .......................................... 29
Growth Hormone ...................................................................................................... 30
Excess of Deficiency of Growth Hormone ............................................................... 35
Growth Hormone and Obesity.................................................................................. 36
Leptin and Growth Hormone .................................................................................... 39
Objectives.................................................................................................................. 41
2 GENERAL METHODS
43
Animals ..................................................................................................................... 43
Diets .......................................................................................................................... 43
Feeding, Pair-feeding, and Body Weight Measurements ......................................... 43
Leptin Challenge Test............................................................................................... 44
Alzet Osmotic Minipumps ........................................................................................ 45
Right Atrial Cannulation........................................................................................... 45
Anesthesia ............................................................................................................. 45
Preparation ............................................................................................................ 45
v
Surgery.................................................................................................................. 46
Recovery ............................................................................................................... 46
Blood sampling cages ............................................................................................... 46
Blood Sampling and Tissue Collection..................................................................... 47
Blood Collection from Cannulae .......................................................................... 47
Blood Collection from Tail Vein .......................................................................... 47
Blood Collection via Cardiac Puncture................................................................. 47
Trunk Blood Collection ........................................................................................ 48
Collection of Hypothalamus ................................................................................. 48
Pituitary and Organ Weights................................................................................. 48
Cell Culture............................................................................................................... 48
GH1 Cells.............................................................................................................. 48
Plating Cells .......................................................................................................... 49
Five-Day Time-Course ......................................................................................... 49
Media Experiment................................................................................................. 49
Collecting RNA from GH1 Cells.......................................................................... 50
Radioimmunoassays.................................................................................................. 50
GH Iodination ....................................................................................................... 50
Growth Hormone RIA .......................................................................................... 51
IGF Iodination....................................................................................................... 51
IGF RIA ................................................................................................................ 52
Leptin RIA ............................................................................................................ 52
Insulin RIA............................................................................................................ 53
Glucose and Triglyceride Assays.............................................................................. 54
DNA Assay ............................................................................................................... 54
RNA extraction, RT-PCR, Southern Blot................................................................. 55
Protein Extraction and Western Immunoblotting ..................................................... 57
Protein Extraction ................................................................................................. 57
Micro BCA Protein Assay .................................................................................... 58
Western Immunoblot Analysis for Leptin Receptor ............................................. 59
3 CIRCULATING GROWTH HORMONE LEVELS ARE ELEVATED IN
RATS FED A HIGH-FAT DIET
62
Introduction............................................................................................................... 62
Methods..................................................................................................................... 63
Animals ................................................................................................................. 63
Cannulation Surgery, Blood Sampling, and Tissue Collection ............................ 64
Radioimmunoassays.............................................................................................. 65
RT-PCR for Leptin Receptor ................................................................................ 65
Statistics .................................................................................................................... 66
Results ....................................................................................................................... 66
Body Weight ......................................................................................................... 66
Food Intake ........................................................................................................... 66
Leptin .................................................................................................................... 67
Leptin Receptor mRNA ........................................................................................ 67
vi
IGF ........................................................................................................................ 67
Growth Hormone Profile ...................................................................................... 67
Discussion ................................................................................................................. 72
4 LEPTIN TREATMENT INCREASES GROWTH HORMONE SECRETION
IN CULTURED GH1 CELLS
77
Introduction............................................................................................................... 77
Methods..................................................................................................................... 79
GH1 Cells.............................................................................................................. 79
Plating Cells and Leptin Concentration................................................................ 79
Five-Day Time-Course ......................................................................................... 79
Media Experiment................................................................................................. 80
RT-PCR for Leptin Receptor ................................................................................ 80
Western Immunoblot for Leptin Receptor ............................................................ 80
GH Radioimmunoassay ........................................................................................ 81
Statistics ................................................................................................................ 81
Results ....................................................................................................................... 81
Ob-Rb mRNA and Protein Expression ................................................................ 81
Five-Day Time-Course ......................................................................................... 82
Media Experiment................................................................................................. 82
Discussion ................................................................................................................. 85
5 LEPTIN RESISTANCE IS ASSOCIATED WITH HYPOTHALAMIC
RECEPTOR mRNA AND PROTEIN DOWNREGULATION
88
Introduction............................................................................................................... 88
Methods..................................................................................................................... 89
Animals ................................................................................................................. 89
Groups................................................................................................................... 90
Serum Measurements ............................................................................................ 90
Leptin Challenge Test Pilot Study........................................................................ 91
Leptin Challenge Test........................................................................................... 91
RT-PCR for Leptin Receptor ................................................................................ 91
Western Immunoblot for Leptin Receptor ............................................................ 92
Statistics .................................................................................................................... 92
Results ....................................................................................................................... 93
Leptin .................................................................................................................... 93
Body Weight ......................................................................................................... 93
Food Intake ........................................................................................................... 93
Leptin Challenge Test Pilot Studies...................................................................... 94
Leptin Challenge Test........................................................................................... 94
Leptin Receptor mRNA in Hypothalamus ............................................................ 94
Leptin Receptor Protein Expression in Hypothalamus ......................................... 94
IGF-1 Values......................................................................................................... 95
Hormonal and Metabolic Measures ...................................................................... 95
vii
Discussion ............................................................................................................... 100
6 LEPTIN'S EFFECTS ON FOOD INTAKE AND BODY WEIGHT ARE
DIFFERENTIALLY ATTENUATED IN RATS FED A LOW-FAT OR HIGHFAT DIET
107
Introduction............................................................................................................. 107
Methods................................................................................................................... 108
Animals and Diets ............................................................................................... 108
Osmotic Pumps, Blood Sampling, and Body Temperature ................................ 109
Leptin and IGF-1 Radioimmunoassays............................................................... 109
Statistics .................................................................................................................. 110
Results ..................................................................................................................... 110
Effects of Diet on Food Intake ............................................................................ 110
Leptin .................................................................................................................. 110
Effects of Diet and Leptin on Food Intake.......................................................... 111
Effects of Diet and Leptin on Body Weight ....................................................... 111
Effects of Diet and Leptin on IGF-1 Values ....................................................... 112
Effects of Diet and Leptin on Organ Weights..................................................... 112
Effects of Diet and Leptin on Body Temperature............................................... 112
Discussion ............................................................................................................... 119
7 GENERAL DISCUSSION
125
REFERENCES
134
BIOGRAPHICAL SKETCH
166
viii
LIST OF FIGURES
Figure
Page
Figure 2-1: MgCl 2 (mM) and temperature (°C) optimization..................................................56
Figure 2-2: Cycle optimization...............................................................................................56
Figure 2-3: Southern blot of leptin receptor ...........................................................................57
Figure 2-4: Representative RT-PCR from hypothalamus samples..........................................57
Figure 2-5: Trizol extraction ..................................................................................................58
Figure 2-6: Cell lysis buffer extraction...................................................................................58
Figure 2-7: Protein loading optimization................................................................................61
Figure 2-8: Primary antibody concentration optimization.......................................................61
Figure 3-1: Body Weight ........................................................................................................68
Figure 3-2: Food Intake ..........................................................................................................69
Figure 3-3: Leptin Levels .......................................................................................................69
Figure 3-4: Ob-Rb mRNA......................................................................................................70
Figure 3-5: IGF.......................................................................................................................70
Figure 3-6: GH Profile ...........................................................................................................71
Figure 4-1: Ob-Rb mRNA on GH1 Cells ...............................................................................82
Figure 4-2: Five-Day Time-Course........................................................................................83
Figure 4-3: Media Supplemented with 10% Horse Serum and 2.5% FBS .............................83
Figure 4-4: Serum-Free Media ...............................................................................................84
Figure 4-5: Media Supplemented with 12.5% Charcoal Stripped FBS..................................84
ix
Figure 5-1: Leptin Levels Indicate Proper Pump Activity ...................................................... 95
Figure 5-2: Body Weight is Dose-Dependently Decreased by Leptin Treatment.................... 96
Figure 5-3: Food Intake is Initially Decreased by Leptin; Ultimately Resistance Develops... 96
Figure 5-4: Leptin Challenge Test Pilot Study........................................................................ 97
Figure 5-5: Leptin Challenge Test.......................................................................................... 97
Figure 5-6: Leptin Receptor mRNA in Hypothalamus ............................................................ 98
Figure 5-7: Leptin Receptor Protein Expression in Hypothalamus ......................................... 98
Figure 5-8: IGF-1 Values ....................................................................................................... 99
Figure 6-1: Absolute Food Intake Reported in Various Diet Parameters................................113
Figure 6-2: Leptin Levels Indicate Proper Pump Activity ......................................................114
Figure 6-3: The Effects of Leptin on Food Intake in Diets of Varying Calorie and Fat
Contents ................................................................................................................115
Figure 6-4: The Effects of Leptin on Food Intake - Normalized to 100 grams Body
Weight ..................................................................................................................116
Figure 6-5: The Effects of Leptin on Body Weight in Diets of Varying Calorie and Fat
Contents ................................................................................................................117
Figure 6-6: IGF-1 Values .......................................................................................................118
x
LIST OF TABLES
Table
Page
Table 1-1: Obesity Models.....................................................................................................
5
Table 1-2: Orexigenic and Anorexigenic Neuropeptides Regulated by Leptin....................... 18
Table 1-3: Leptin Receptors................................................................................................... 23
Table 1-4: Mediators that Inhibit GH Secretion in Both Man and Rodents ............................ 32
Table 1-5: Mediators that Stimulate GH Secretion in Both Man and Rodents........................ 32
Table 3-1: Growth Hormone Profile of Diet-Treated and Control Rats ................................. 71
Table 5-1: Hormonal and Metabolic Measures...................................................................... 99
Table 6-1: Organ Weights ......................................................................................................118
Table 6-2: Body Temperature ................................................................................................119
Table 7-1: Model of GH Regulation by Leptin.......................................................................133
xi
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
LEPTIN RESISTANCE REDUCES GROWTH
HORMONE SECRETION AND CONTRIBUTES TO
THE PATHOGENESIS OF OBESITY
By
Robin Leigh Martin
May 2000
Chairman: William J. Millard
Major Department: Pharmacodynamics
Leptin is secreted by adipocytes in amounts indicative of body fat content.
Variations in leptin levels contribute to hypothalamic regulation of feeding and
metabolism to maintain body weight homeostasis. Importantly, if there is increasing
leptin with increasing body fat and the hypothalamus cannot detect or respond to it, such
as is seen in leptin resistance, obesity may develop. Growth hormone (GH) is secreted by
the anterior pituitary and directly influences body composition through lipolysis. GH
also stimulates the secretion of insulin-like growth factor I (IGF-1), which mediates some
of the actions of GH.
The hypotheses of this dissertation are (1) under normal circumstances, leptin
stimulates GH, and (2) in the leptin resistance situation described above, leptin fails to
xii
stimulate GH. In obesity, GH is severely attenuated, and leptin resistance may be the
mechanism by which this occurs.
To test the first hypothesis, rats were given diets with varying fat contents and GH
was measured. The diet with the highest fat content did not cause obesity; there were
elevated levels of leptin but not enough to produce resistance. These animals had an
enhanced GH-IGF-1-axis. In a second study, leptin treatment resulted in elevated GH
secretion from a GH-secreting cell line (GH1), further strengthening the hypothesis.
To test the second hypothesis, rats were given normal rat chow and implanted
with osmotic minipumps for the continuous infusion of two doses of leptin or vehicle.
The leptin-treated animals developed resistance, as measured by loss of effect on food
intake but not on various metabolic measures, including glucose, triglycerides, and
insulin. IGF-1 was attenuated in the high-dose leptin group. Leptin receptors were
reduced in this study. A final study utilized both the diets of varying fat content and the
leptin-filled osmotic minipumps. Leptin was shown to lose its effectiveness in animals
fed the high-fat diet and IGF-1 was attenuated. The rats were fed the diets for a longer
period than in the first diet study, allowing time for the development of leptin resistance.
IGF-1 was also attenuated in animals infused with leptin. The results of these two studies
agree with the second hypothesis.
CHAPTER 1
LITERATURE REVIEW
Obesity
Obesity is one of the most common health problems in industrial societies
[Grundy and Barnett 1990]. Obesity is a health threat in part because it is associated with
many other diseases, including type II diabetes, gallstones, and certain cancers [Grundy
and Barnett 1990] and with excessive mortality [NIH 1985]. Data from National Health
and Nutrition Examination Surveys (NHANES) show a high correlation between obesity
and risk for coronary artery disease [NIH 1985]. It has also been shown that most obese
people experience hypertriglyceridemia and hypercholesterolemia [Grundy and Barnett
1990]. In addition, obesity is a very strong risk factor for hypertension. Obesity is
epidemic in the United States (US) [Wilding 1998] and its prevalence is increasing
[Grundy and Barnett 1990].
Currently, the criteria for inclusion into the overweight and obese categories are a
relationship of height and weight [Bray 1992b]. Body mass index (BMI) is body weight
in kilograms divided by height in meters squared. In adults aged 19-34 years, BMI of 1925 kg/m2 is considered normal, 25-30 kg/m2 is overweight, and >30 kg/m2 constitutes
obesity. In people 35 years and older, the scale for normal weight is BMI of 19-27
kg/m2, 27-30 kg/m2 is considered overweight, and >30 kg/m2 represents obese. In the US
population of adults aged 25 years or more, 42% of men and 28% of women are
overweight and 21% of men and 27% of women are obese [Must et al. 1999]. In 1991,
1
2
the prevalence of obesity was 12.0% [Mokdad et al. 1999] and increased to 17.9% in
1998. The incidence of obesity in every state, in both sexes, and in all age groups, races,
and educational levels was increased. Nearly 10% of total health care costs are related to
obesity [Wilding 1998].
We often blame excessive eating and insufficient exercise for obesity; however
there is still no concrete explanation for the fact that some people become obese, despite
attempts not to, and others, apparently without much effort, do not [Ravussin and
Danforth 1999]. Genetic contributions to the development and maintenance of obesity
cannot be eliminated. Studies in twins demonstrated an inherited tendency to gain weight
in response to overfeeding [Wilding 1998]. Additionally, it was recently shown that
there is unaccounted for physical activity in some people that prevents weight gain
[Levine et al. 1999]. This activity has been called NEAT, for nonexercise activity
thermogenesis, and is made up of "nonvolitional muscle activity" including maintenance
of posture, fidgeting, and muscle tone. In some people, NEAT is triggered by excess
food and prevents fat gain. This concept was proposed in the past [Widdowson et al.
1954] with the suggestion that fidgetiness was more important in prevention of weight
gain than seemed obvious. It was shown that increased fidgeting enhanced energy
expenditure [Rassuvin et al. 1986; Zurlo et al. 1992]. This may account for some of the
individual variations observed in food consumption, exercise, and body weight
fluctuations.
Although obesity is strongly believed to be a risk factor in many diseases, some
experts are not in agreement. One professor suggests that obesity is simply a
consequence of a lifestyle, such as physical inactivity, that is associated with elevated
3
risks for certain disease, and that obesity in itself is not a risk factor [McDonald 1996].
There are some supporters of this theory, however obesity is intertwined with lack of
physical activity and most researchers are in agreement that obesity poses health risks. It
has been suggested that, without a change in the progression of obesity in today's society,
there will be increasingly overwhelming health care costs and hazardous health
conditions related to this disease [Must et al. 1999]. Thus, research relating to obesity is
important on an individual as well as on a national level.
Obesity Models
There are many types of genetic mutations that result in pathophysiological
signaling and metabolic alterations and which cause obesity in rodents [Guerre-Millo
1997]. Two such models are the obese (ob/ob) and diabetes (db/db) mice, both of which
exhibit hyperphagia, profound early-onset obesity, hyperglycemia, hyperinsulinemia,
infertility [Coleman 1973], and defective thermoregulation [Coleman 1978] indicative of
a hypothalamic defect. The phenotypes of the ob/ob and db/db mice are identical and the
strains can only be distinguished by genetic mapping or through parabiosis studies
[Coleman 1973; Coleman 1978]. To study these strains of mice, parabiosis studies were
completed in the following pairs of mice: db/db + ob/ob, ob/ob + normal, and db/db +
normal. Parabiosis is the surgical joining of two mice in a manner in which they share
the same circulatory system. In the db/db + ob/ob parabiosis study, the ob/ob partner
became hypoglycemic, lost weight, and starved to death [Coleman 1973]. A similar
phenomenon was seen in the normal partner in the parabiosis experiment joining db/db +
normal mice [Coleman 1969]. The results of these experiments suggest that there are
satiety centers in the ob/ob and normal mice that respond to a circulation factor that is
4
produced by db/db mice. The db/db mice do not respond to the circulating satiety factor,
suggesting that the target satiety center is defective [Coleman 1973]. When ob/ob mice
were joined with normal mice, there was no alteration in the feeding behavior of the
normal mice suggesting that the ob/ob mice do not make the circulating satiety factor in
sufficient quantities to affect behavior [Coleman 1973].
The ob/ob genetic defect in mice was first discovered in the laboratory of Snell in
1950 [Ingalls et al. 1950]. Ob/ob is the result of an autosomal recessive mutation
[Coleman 1978] of the obese gene located on chromosome 6, which is also the location
of the leptin gene [Zhang et al. 1994] (leptin will be discussed in detail later in the
chapter). There are two different ob/ob strains of mouse: one with an absence of ob
mRNA [Zhang et al. 1994], and the other with 20 times higher expression of a mutant
protein [Zhang et al. 1994; Frederich et al. 1995b]. That leptin mRNA is altered in this
mutant suggests that the ob gene can be regulated transcriptionally [Mason et al. 1998].
The phenotype of the mice used in this study includes obesity, hyperphagia, low oxygen
consumption and body temperature, and reduced physical activity.
The db/db genetic defect in mice was first discovered in the laboratory of
Coleman in 1966 [Coleman and Hummel 1966; Hummel et al. 1996]. Db/db is the result
of an autosomal recessive mutation [Coleman 1978] of the diabetes gene located on
chromosome 4, which is the location of the leptin receptor gene [Tartaglia et al. 1995].
The db phenotype results from a mutation of the intracellular portion of the leptin
receptor that normally initiates signal transduction [Flier and Elmquist 1997] and as such
affects only the long form of the receptor [Guerre-Millo 1997]. Leptin is able to bind
5
specifically and with high affinity to the choroid plexus in db/db mice [Devos et al.
1996], indicating that the short form of the leptin receptor is present.
The fa/fa genetic defect in rats was first discovered in the laboratory of Zucker
and Zucker in 1961 as an autosomal recessive mutation mapped to chromosome 5 [Truett
et al. 1991]. The fa mutation is in the same gene as the db mutation [Chua et al. 1996].
The fa mutation is a missense mutation in the extracellular domain of the leptin receptor
[Chua et al. 1996; Flier and Elmquist 1997] that results in inadequate transport to the
plasma membrane [Flier and Elmquist 1997]. As in db/db mice, leptin is able to bind
specifically and with high affinity to the choroid plexus in fa/fa rats [Devos et al. 1996].
The Koletsky strain of rat develops obesity, hyperinsulinemia, hypertension, and
proteinuria [Koletsky et al. 1973] due to a single recessive gene with a mutation in the
extracellular domain of the leptin receptor [Takaya et al. 1996]. The same gene as is
mutated in the Zucker fa/fa rat is mutated in the Koletsky rat, however, the Koletsky
mutation results in a premature stop codon, making this strain of rat a leptin receptor null
model.
Table 1-1: Obesity Models
Model
Chromosome
Ob/ob mouse
6
Db/db mouse
4
Fa/fa rat
5
Koletsky rat
5
Anomaly
Absence of leptin
mRNA or expression of
mutant protein
Mutation of intracellular
portion of leptin
receptor (long form)
Missense mutation of
extracellular portion of
leptin receptor
Premature stop codon –
leptin receptor null
Reference
Ingalls et al. 1950
Coleman and Hummel 1966
Hummel et al. 1966
Zucker and Zucker 1961
Koletsky et al. 1973
6
In addition to genetically occurring models of obesity, obesity models can be
created by chemical damage to the hypothalamus. For example, the monosodium
glutamate (MSG)-treated animal develops a syndrome in which obesity is its most
characteristic feature [Olney 1969]. MSG treatment results in lesions of the brain,
specific to those areas surrounding the third ventricle such as the arcuate nucleus of the
hypothalamus. Females are infertile and have atrophied ovaries, uteri, and endometria;
males can reproduce and have normal testes. MSG rodents gain weight, which is more
evident in females, without accompanying hyperphagia, in fact, they appear to be slightly
hypophagic [Olney 1969]. The obesity in this model results from metabolic disturbances,
which is partially demonstrated in their decrease in body length [Olney 1969]. This
decrease in length suggests that there is a defect in growth hormone production,
secretion, and/or activity. MSG-treated rats are also lethargic as adults. As a result of
damage to the arcuate nucleus, the MSG-treated rodent has elevated leptin mRNA and
protein expression compared to control [Frederich et al. 1995b].
Theories of Food Intake
Several theories have been proposed to describe the mechanism of initiation of
food consumption. Most of the theories revolve around the dual center hypothesis
proposed in 1951 by Anand and Brobeck. The dual center hypothesis suggested that the
lateral region of the hypothalamus, the "feeding center," played a role in the initiation of
feeding and the ventromedial region, the "satiety center," inhibited feeding. This
hypothesis was based on studies that showed that lesions to the ventromedial
hypothalamus (VMH) and surrounding areas resulted in hyperphagia and obesity
[Hetherington et al. 1940] and lesions to the lateral hypothalamus (LH) abolished food
7
intake [Anand and Brobeck 1951]. More recent studies have shown the dual center
hypothesis to be too simplistic to be entirely accurate. When lesions of the VMH were
restricted solely to that nucleus of the hypothalamus, hyperphagia and obesity did not
develop [Reynolds 1963]. In addition, it was shown that hyperphagia developed
following injury to other nuclei of the hypothalamus [Jansen and Hutchinson 1969],
including areas through which critical catecholamine tracts travel [Sclafani and Berner
1977]. Another study demonstrated that obesity could develop in the absence of
hyperphagia [Han 1968]. Although these studies argue against the basic dual center
hypothesis, they demonstrate the importance of the hypothalamus in food intake and the
development of obesity [Bray and York 1979].
The aminostatic theory of food intake was proposed by Rogers and Leung in
1973. The theory suggests that rats eat less when fed a diet devoid of certain amino
acids, a diet with an amino acid imbalance, or a diet high in protein. The authors
suggested that brain amino acid content closely resembled the amino acid content in the
blood, and that a receptor system, which recognizes blood amino acid concentration, is
located in the brain. Their results point to a central involvement in orexic behavior.
The glucostatic theory was proposed by Mayer in 1953. The hypothesis was that
there are glucose-sensitive receptors, glucoreceptors, in the lateral hypothalamus that
initiate feeding when blood glucose is low. Additionally, the hypothesis suggests that
there are glucoreceptors in the satiety center of the hypothalamus that terminate feeding
when activated by elevated blood glucose. It was discovered that the mere presence of
glucose was not sufficient to initiate feeding, but that glucose had to cross the membranes
of the glucose-sensitive cells or undergo oxidation. It was later shown that a wide range
8
of metabolites in the blood in addition to glucose are capable of initiating or terminating
feeding.
The thermostatic theory of food intake was developed by Brobeck in the late
1940s [Brobeck 1948]. The theory states that "animals eat to keep warm and stop eating
to prevent hyperthermia." When animals are in warmer temperatures, they will decrease
their intake of food to prevent the production of too much heat within the body.
Consequently, when animals are in cooler temperatures, food intake increases in an effort
to produce heat and prevent hypothermia. It was further hypothesized that the amount of
food ingested is dependent on how much heat the food produces and how much heat is
required to maintain body temperature. Kennedy argued against this theory [1953]
because he felt that the weight loss experienced by these animals was due to the elevated
temperatures to which the animals were exposed which may have resulted in dehydration.
Kennedy developed his own theory of food intake in 1950 referred to as the
lipostatic theory. This theory is similar to Mayer's theory and agrees with the role of the
hypothalamus, but utilizes a signal other than glucose. It was theorized that, since young
rats are so adept at maintaining constancy of fat stores, there must be some circulating
factor that, along with the hypothalamic mechanism, regulates feeding and energy
expenditure. The circulating factor was proposed to be some unknown factor of the
synthesis, transport, or metabolism of fat. In 1994, a circulating peptide hormone
synthesized in fat was discovered [Zhang et al. 1994] which fit the role of Kennedy’s
lipostatic factor. This hormone is leptin.
9
Leptin
Leptin, coming from the Greek word leptos, meaning thin, is a hydrophobic, 167amino acid, 16-kilodalton protein hormone [Samson et al. 1996] transcribed by the ob
gene. Leptin is often referred to as the obese protein. The 4.5 kilobase mRNA from
which leptin is translated is 84% homologous between humans and mice and has 67%
conservation among a large variety of vertebrates, including human, gorilla, chimpanzee,
orangutan, rhesus monkey, dog, cow, pig, rat, and mouse [Zhang et al. 1997]. McGregor
et al. [1996] demonstrated that mouse leptin has no homology with other known proteins.
Leptin has features of a protein that is secreted [Zhang et al. 1994; Cohen et al.
1996b]. It contains a disulfide bond in the carboxy-terminus [Cohen et al. 1996b]; a
mutation of either of the conserved cysteine residues that form the disulfide bond results
in the loss of activity [Zhang et al. 1997]. The fragment of leptin that is the predicted
signal peptide is 1-21 and fragment 22-56 may be the portion of the protein that is
biologically active [Samson et al. 1996]. Additionally, domains between residues 106140 have been shown to induce satiety [Grasso et al. 1997].
Leptin is produced in and secreted mainly from mature white adipocytes. The
other types of fat cells (preadipocytes, young adipocytes, fibroblasts, endothelial cells,
Schwann cells, and vascular cells) do not express the ob gene [Frederich et al. 1995b;
Maffei et al. 1995a]. In culture, leptin mRNA is not expressed until mouse fibroblasts
are fully differentiated into adipocytes [Yoshida et al. 1996]. Furthermore, leptin is
produced in fat depots throughout the body, including epididymal, parametrial,
abdominal, perirenal, and inguinal [Maffei et al. 1995a; McGregor et al. 1996]. Leptin is
also expressed in brown adipose tissue [Maffei et al. 1995a], but to a much lesser extent
than in white adipose tissue. In addition to expression in adipose tissue, leptin protein or
10
its mRNA have been observed in placenta [Senaris et al. 1997], amniotic fluid
[Schubring et al. 1997], and cord blood [Matsuda et al. 1997; Schubring et al. 1997].
Recently, leptin mRNA has also been found in stomach [Bado et al. 1998], skeletal
muscle [Wang et al. 1998c], brain [Esler et al. 1998; Wiesner et al. 1999], and pituitary
[Jin et al. 1999; Morash et al. 1999; Jin et al. 2000].
Leptin is secreted in proportion to adipose mass [Maffei et al. 1995b; Considine et
al. 1996b] and consistent with fat cell size; therefore, heavier humans and animals
generally produce more leptin than lean members of the same species do. Leptin is not
stored in adipocytes [Rau et al. 1999].
Leptin is secreted in a pulsatile manner with short pulses lasting just over 30
minutes [Licinio et al. 1997]. It has been reported that leptin secretion demonstrates
circadian and ultradian rhythms similar to other endocrine hormones [Sinha et al. 1996a;
Sinha et al. 1996c]. Leptin's circadian rhythm exhibits increased pulse frequency at night
[Sinha et al. 1996a; Licinio et al. 1997] and decreased activity in the afternoon [Sinha et
al. 1996a] in both lean and obese humans. Perhaps the purpose of increased leptin at
night is to suppress appetite during sleep [Sinha et al. 1996a]. It is interesting to note that
this diurnal pattern of leptin secretion is the inverse of that of adrenocorticotropin
hormone (ACTH) and cortisol [Licino 1997], however, the apparent diurnal rhythm of
leptin seems to be more related to feeding than to time of day [Saladin et al. 1995].
Perhaps leptin helps regulate the diurnal pattern of glucocorticoids [Ahima et al. 1996].
It was suggested that, like other hormones, this pulsatile secretion of leptin might
be required for maximum effectiveness [Licino 1997]. Leptin-producing fat depots,
however, do not resemble other endocrine glands; they are of varying sizes and are
11
located throughout the body, and as such, would be difficult to coordinate into producing
a circadian secretion pattern [Sinha et al. 1996c]. It has been suggested that the apparent
pulsatility displayed by leptin actually comes from regulation of its clearance or
elimination [Licino 1997].
Upon secretion, leptin distributes throughout the extracellular space and
accumulates in cellular water [Cumin et al. 1996] or circulates in the blood either free or
complexed with binding proteins [Houseknecht et al. 1996]. Several circulating leptinbinding proteins and a soluble leptin receptor have been observed [Lee et al. 1996]. In
lean humans, the majority of circulating leptin is bound [Sinha et al. 1996b]. Perhaps the
purpose of leptin binding is to limit bioavailability and regulate its actions on feeding and
metabolism homeostasis [Sinha 1997]. Binding proteins also protect leptin from
premature degradation or elimination [Liu et al. 1997].
Leptin that is bound remains in circulation and has a half-life of 71 minutes [Hill
et al. 1998]. Free leptin is rapidly degraded by proteases and has a half-life of less than 4
minutes [Sharma et al. 1997]. In patients with chronic renal failure [Iida et al. 1996] and
end-stage renal disease [Merobet et al. 1997] and in chronic hemodialysis patients
[Sharma et al. 1997] leptin levels are elevated, largely represented by the pool of free
leptin [Sharma et al. 1997]. In addition, the molecular weight (16-kDa) and half-life
(approximately 25 minutes) [Klein et al. 1996] of leptin are similar to other peptide
hormones that are degraded by the proximal tubules [Bennett and McMartin 1979],
indicating that the kidneys are capable of metabolizing leptin [Klein et al. 1996]. Leptin
receptors have been found in the kidneys [Tartaglia et al. 1995] and leptin has, in fact,
been shown to have a renal mechanism of elimination [Cumin et al. 1996; Iida et al.
12
1996; Merabet et al. 1997; Sharma et al. 1997]. It was found that leptin is cleared in a
two-pool model [Hill et al. 1998] with approximately 75% of leptin being cleared within
the first few minutes of secretion [Cumin et al. 1996].
These results, taken together, strongly indicate that leptin is degraded and/or
filtered by the kidneys. It is important to note that when pharmacological doses of leptin
are administered, the pathways involved in elimination do not become saturated [Cumin
et al. 1996; Hill et al. 1998]. Therefore, neither leptin distribution nor half-life differs
between lean and fat rats [Vila et al. 1998]. Furthermore, it was shown that
malfunctioning clearance or turnover rates [Klein et al. 1996] cannot account for the
hyperleptinemic characteristic of obesity.
Roles of Leptin
Leptin has been implicated as the lipostatic signal that is sensed, directly or
indirectly, by the hypothalamus [Campfield et al. 1995; Maffei et al. 1995a; Stephens et
al. 1995; Ahima et al. 1996; Levin et al. 1996; Schwartz et al. 1996a; Vaisse et al. 1996;
Woods and Stock 1996; Dawson et al. 1997]. Leptin in transported across the bloodbrain barrier by a saturable process [Banks et al. 1996] making it possible that this be the
rate-limiting step in leptin bioactivity [Banks et al. 1996; Schwartz et al. 1996b; Golden
et al. 1997]. In the hypothalamus, leptin works specifically in nuclei responsible for
feeding homeostasis: arcuate, ventromedial, paraventricular, lateral, and ventral
premammillary [Mercer et al. 1996]. Leptin is also active in areas of the brain which
may be important in controlling metabolism, energy balance, and transport into the brain
[Steiner 1996] such as cerebellum, choroid plexus, leptomeninges, cortex, hippocampus,
and thalamus [Mercer et al. 1996; Steiner 1996].
13
Leptin acts to regulate body weight and the size of adipose depots by the
regulation of food intake and energy expenditure [Zhang et al. 1994; Campfield et al.
1995; Halaas et al. 1995; Pelleymounter et al. 1995; Halaas et al. 1997]. It has been
suggested that leptin is an afferent signal in a negative feedback loop between adipose
tissue and the appetite/satiety centers in the brain [Rohner-Jeanrenaud and Jeanrenaud
1996]. Exogenous leptin administration is effective at reducing food intake in animals of
normal body weight [Campfield et al. 1995; Halaas et al. 1995] as well as in animals with
obesity due to defective leptin production [Pelleymounter et al. 1995]. Daily
intraperitoneal leptin injections given to ob/ob mice result in a dose- and time-dependent
reduction in food intake and body weight [Pelleymounter et al. 1995]. The effects of
leptin on food intake are lost over time, but the effects of leptin on body weight are not.
Leptin treatment also restores the oxygen consumption, body temperature, and activity
levels in these leptin-deficient mice to levels seen in control wild type mice. When leptin
treatment is terminated, body weight levels return to those of control animals [Campfield
et al. 1995]. Leptin treatment is also effective when administered continuously via
osmotic pumps or when administered centrally [Campfield et al. 1995].
It has been shown that leptin-treated animals lose body weight even when pair-fed
the same amount, caloric content, and fat content of food as that consumed by control
animals. Factors other than those involved with feeding are implicated in the control of
body weight. The rate of metabolism, for example, makes a considerable difference in
body composition among individuals, and, as previously mentioned, leptin increases
metabolism [Zhang et al. 1994; Campfield et al. 1995; Halaas et al. 1995; Pelleymounter
et al. 1995; Halaas et al. 1997].
14
Brown adipose tissue (BAT) in small mammals is important in heat production
and metabolism. When BAT cells are stimulated, mitochondria express uncoupling
protein 1 (UCP1) [Cinti 1992]. The sympathetic nervous system activates existing as
well as stimulates synthesis of new UCP1 [Zhao et al. 1994]. Upon activation, UCP1
uncouples mitochondria, which results in elevated substrate oxidation [Klingenberg
1990] and thermogenesis. It has been shown that leptin increases energy expenditure and
oxygen consumption in rats [Scarpace et al. 1997]; the mechanism by which this occurs
is an increase in UCP1 mRNA [Scarpace et al. 1997] via sympathetic activation
[Scarpace and Matheny 1998]. In addition, leptin stimulates sympathetic outflow to BAT
[Collins et al. 1997].
It has been shown that uncoupling protein 2 (UCP2) can also uncouple
mitochondria, and acts in white adipose tissue (WAT) as well as in BAT [Fleury et al.
1997; Scarpace and Matheny 1998]. The mechanism by which leptin stimulates UCP2
does not require sympathetic activation, but rather utilizes some as yet unexplained
indirect mechanism [Commins et al. 1999]. Together, these results indicate that leptin
activates the sympathetic nervous system and stimulates the production of UCP both
directly and by hypothalamic mechanisms, resulting in increased thermogenesis.
By reducing feeding and increasing metabolism, leptin decreases fat content.
Leptin also reduces fat by other means. Central leptin treatment degrades adipocytespecific genomic DNA in a ladder pattern consistent with apoptosis [Qian et al. 1998].
There is also evidence of condensed chromatin and histological staining consistent with
apoptotic events. In addition, leptin inhibits acetyl-CoA carboxylase (ACC) activity [Bai
15
et al. 1996]. ACC is the enzyme that represents the rate-limiting step in fatty acid
synthesis. By inhibiting ACC activity, leptin inhibits lipogenesis.
Both synthesis and degradation of fat regulate energy stores. In addition to
inhibiting lipogenesis, leptin triggers lipolysis by enhancing mitochondrial fatty acid
oxidation. This oxidation results in attenuated intracellular fatty acid and triglyceride
concentrations [Bai et al. 1996; Qian et al. 1998]. Leptin administration causes weight
loss by reduction of fat mass [Halaas et al. 1995] whereas food deprivation alone causes
weight loss by reduction of both fat and lean mass.
It is clear that leptin is the protein that, consistent with the lipostatic theory of
food intake, acts as a homeostatic marker of adipose tissue that regulates the size of
adipose mass [Zhang et al. 1994; Frederich et al. 1995b]. Elevated leptin is considered to
be a marker of the obese state [Maffei et al. 1995b], but it is also important in the
prevention of starvation [Ahima et al. 1996; Spiegelman and Flier 1996; Flier and
Elmquist 1997]. Evolutionarily, the role of leptin in starvation may be more important
than its role in the prevention of obesity. Leptin is also involved in cardiovascular, renal,
and reproductive physiology, and may be part of the circuitry of reward pathways.
Regulation of Leptin
It has been observed that the downregulation of leptin mRNA expression is
dependent on physiologically active leptin receptors [Guerre-Millo 1997]. If the receptor
is mutated or otherwise inactive, the result will be overexpression of leptin. Leptin
mRNA levels may be regulated by tissue-specific transcription factors or by endocrine
and/or paracrine hormonal regulators [Mason et al. 1998]. Regulatory regions of the
leptin gene promoter include sites specific for transcription factors that control adipocyte
16
differentiation [Auwerx and Staels 1998]. CCAAT/enhancer-binding protein α
(C/EPBα) induces leptin gene expression and peroxisome proliferator-activated receptorγ (PPARγ) inhibits leptin gene expression [Hollenberg et al. 1997].
Leptin levels are acutely altered in response to feeding [Caro et al. 1996b].
Leptin levels are low in food-deprived normal [MacDougald et al. 1995; Hardie et al.
1996; Sinha et al. 1996b] and db/db mice [Frederich et al. 1995b] and rise upon
refeeding. Leptin levels can be restored, with leptin treatment, to levels found in fed
mice [Ahima et al. 1996]. Human subjects fasted for 24 hours may lose only 0.5% of
body fat but 50% of leptin concentration [Boden et al. 1996]. It is clear that leptin
expression is regulated in response to nutritional alterations that influence adipose mass
[Frederich et al. 1995b].
Leptin and Hormonal Interactions
Leptin is intricately intertwined with many central and peripheral hormones. In
the brain, leptin affects many neuropeptide systems responsible for the regulation of
feeding. Neuropeptide Y (NPY) [Kalra et al. 1989], β-endorphin [McKay et al. 1981],
agouti [Lu et al. 1994], agouti-related peptide (AgRP) [Ollmann et al. 1997; Shutter et al.
1997], melanin concentrating hormone (MCH) [Qu et al. 1996], galanin [Sahu 1998], and
orexins [Sakurai et al. 1998] stimulate feeding. In general, leptin inhibits peptides that
stimulate feeding. Leptin binds to its receptors in the arcuate nucleus and decreases NPY
mRNA [Stephens et al. 1995; Schwartz et al. 1996c] thereby attenuating the feeding
response normally elicited by NPY. Both agouti [Lu et al. 1994] and AgRP [Ollmann et
al. 1997; Shutter et al. 1997] increase feeding by antagonizing a peptide that inhibits
feeding, α-melanocyte-stimulating hormone (α-MSH). Agouti and AgRP are decreased
17
by leptin [Mizuno and Mobbs 1999; Wilson et al. 1999]. Galanin [Sahu 1998] and
orexins, which act in the lateral hypothalamus to stimulate food intake [Sakurai et al.
1998] are inhibited by leptin [Beck and Richy 1999].
All orexigenic peptides discussed so far are inhibited by leptin. In some cases,
however, leptin’s role in the regulation of these peptides is not as clear. Leptin has been
shown to increase [Huang et al. 1990] as well as decrease [Sahu 1998] MCH. In
addition, leptin receptor mRNA is found on proopiomelanocortin (POMC) mRNAcontaining neurons in the arcuate nucleus [Cheung et al. 1997]. POMC neurons are the
precursors of both stimulatory (β-endorphin) and inhibitory (α-MSH) feeding peptides.
Leptin has been shown both to decrease POMC mRNA [Schwartz et al. 1997; Thornton
et al. 1997] and to increase POMC mRNA [Sahu 1998]. The differential effects of leptin
on POMC mRNA may be the result of differential processing of POMC to β-endorphin
and α-MSH, respectively.
In addition to α-MSH, corticotropin-releasing factor (CRF) [Britton et al. 1982],
cocaine and amphetamine-regulating transcript (CART) [Elias et al. 1998; Kristensen et
al. 1998], and neurotensin [Sahu 1998] inhibit feeding. Just as leptin tends to inhibit
peptides that stimulate feeding, leptin stimulates peptides that inhibit feeding. Leptin
increases CRF mRNA in the paraventricular nucleus of the hypothalamus, potentiating
the action of CRF on the inhibition of feeding [Schwartz et al. 1996c]. CART, which
also is inhibitory on food intake [Elias et al. 1998; Kristensen et al. 1998], is enhanced by
leptin [Elias et al. 1998; Kristensen et al. 1998]. Neurotensin is another hypothalamic
peptide that inhibits feeding behavior, and leptin increases its gene expression [Sahu
1998].
18
Table 1-2: Orexigenic and Anorexigenic Neuropeptides Regulated by Leptin
Orexigenic
Anorexigenic
NPY
β-endorphin
Agouti
AgRP
CRF
MCH
Galanin
Neurotensin
Orexins
α-MSH
In addition to regulation of central peptides, leptin interacts with peripheral
peptides associated with food intake. Blood glucose levels increase directly following a
meal and level off between meals. According to the glucostatic theory of food intake
[Mayer 1953], glucoreceptors in the lateral hypothalamus initiate feeding when blood
glucose is low. Additionally there are glucoreceptors in the satiety center of the
hypothalamus that terminate feeding when activated by elevated blood glucose. During
food deprivation, glucose levels as well as leptin levels are low. Upon ingestion of a
meal, glucose is available to be taken up into cells, and leptin secretion is restored. It has
been shown that glucose administration enhances leptin mRNA in mice [Mizuno et al.
1996] and that small glucose infusions following food deprivation prevent the fall of
leptin levels in humans [Boden et al. 1996].
Insulin is secreted in response to elevations in blood glucose following a meal. In
addition to leptin, insulin would be a potential candidate for the lipostatic theory of food
intake [Kennedy 1950; Kennedy 1953] except for one crucial criterion: it is not produced
in adipocytes. It is generally well agreed-upon that insulin increases leptin mRNA
expression [Yoshida et al. 1996] and leptin production and secretion [Saladin et al. 1995;
Mizuno et al. 1996; Wabitisch et al. 1996; Ahren et al. 1997; Barr et al. 1997], but the
effects of leptin on insulin are less well defined. There is some controversy about
19
whether leptin increases [Barzilai et al. 1997; Sivitz et al. 1997; Tanizawa et al. 1997],
decreases [Cohen et al. 1996a ; Kieffer et al. 1996; Dawson et al. 1997; Poitout et al.
1998], or has no effect [Sinha et al. 1996b] on insulin production, secretion, and/or
action. Perhaps these differences in results lie in the fact that leptin and insulin are both
dynamically regulated by metabolic factors such as meal ingestion and fasting, and both
have an important resistance component, making these comparisons difficult to interpret.
Leptin also has multiple hormonal interactions that may or may not be related to
orexic behavior. Synthesis and secretion of leptin are increased by glucocorticoids [De
Vos et al. 1995; Murakami et al. 1995; Berneis et al. 1996; Slieker et al. 1996; Wabitsch
et al. 1996] and leptin has been proposed as an important feedback regulator of the
hypothalamic-pituitary-adrenal (HPA) axis [Heiman et al. 1997; Licinio et al. 1997;
Pralong et al. 1998]. There is also regulation of leptin by thyroid hormones [EscobarMorreale et al. 1997], growth hormone (GH) [Florkowski et al. 1996], and insulin-like
growth factor-1 (IGF-1) [Bianda et al. 1997]. In turn, leptin is required for maximal
blood GH levels [Carro et al. 1997]. In fasted mice, estrus is delayed, serum testosterone,
luteinizing hormone, and thyroxine levels are reduced, and corticosterone and ACTH are
elevated, all of which can be normalized or nearly normalized with leptin treatment
[Ahima et al. 1996].
Leptin Receptors
Tartaglia et al. [1995] first cloned a high-affinity, single membrane-spanning
receptor for leptin mapped to a gene on chromosome 4 with strong sequence homology
between mouse and human. The receptor was 894 amino acids in length and the
membrane-spanning domain consisted of 23 amino acids. The short intracellular domain
20
of 34 amino acids contained a sequence that allowed binding to Janus protein kinases
(JAKs). It was predicted that the receptor was a member of the class I cytokine receptor
superfamily and was most closely related to gp130 signaling component of the
interleukin-6 (IL-6) receptor, the granulocyte colony-stimulating factor (G-CSF) receptor,
and the leukemia inhibitory factor (LIF) receptor [Tartaglia et al. 1995]. Representatives
of the class I cytokine family to which the leptin receptor has the highest homology have
longer intracellular domains, necessary for signaling, than initially reported by Tartaglia
et al. for the leptin receptor. It was later found that the leptin receptor gene creates a
splice variant with a longer intracellular domain [Lee et al. 1996] capable of signaling.
The gene that encodes the leptin receptor produces multiple splice variants in
varying levels in a tissue-specific manner [Lee et al. 1996]. Initially, four membranebound single-gene splice variants of the receptor were found in the mouse: three (Ob-Ra,
Ob-Rc, and Ob-Rd) with short intracellular domains originally thought to be incapable of
signaling, and one (Ob-Rb) with a longer intracellular domain, through which leptin
exerts its biological effects [Lee et al. 1996]. There is also one soluble leptin receptor
(Ob-Re). More recently, a new form of the receptor, Ob-Rf, was discovered [Wang et al.
1996]. Ob-Rf has strong sequence homology to the previously cloned isoforms, but with
the short intracellular domain dissimilar to the other short forms.
Leptin receptors are located in many areas of the body including lung, kidney,
liver, ovary, testis, prostate, gastrointestinal tract [Cioffi et al. 1996; Lee et al. 1996],
pancreas [Tanizawa et al. 1997] and the central nervous system (CNS). In the CNS,
receptors are especially abundant in nuclei of the hypothalamus implicated in feeding
homeostasis: arcuate, ventromedial, paraventricular, and ventral premammillary [Mercer
21
et al. 1996]. Leptin receptors have also been observed in cerebellum, choroid plexus,
leptomeninges, cortex, hippocampus, thalamus [Mercer et al. 1996; Steiner 1996]. These
areas are not necessarily important in modulating feeding behavior, but may be important
in controlling metabolism, energy balance, and transport into the brain [Steiner 1996].
Leptin binds with high affinity to the choroid plexus and leptomeninges in rats
[Devos et al. 1996]. Ob-Ra is expressed in the choroid plexus (blood-cerebrospinal fluid
or blood-CSF barrier), leptomeninges, brain, hypothalamus, testes, adipose tissue [Lee et
al. 1996], liver, stomach, kidney, heart, lung [Wang et al. 1996] and in rat brain
microvessels that make up the blood-brain barrier [Bjorbaek et al. 1998]. Ob-Ra is
thought to help transport leptin across the blood-brain barrier via receptor-mediated
transcytosis at the microvessels. It was shown that Ob-Ra on human brain endothelium
bind leptin and internalizes it in a temperature-dependent process [Golden et al. 1997].
Leptin may also cross the blood-CSF barrier at the choroid plexus, although this is not the
major means by which leptin gains access to the CSF [Bjorbaek et al. 1998]. It has also
been suggested that Ob-Ra on the leptomeninges degrades CSF leptin [Bjorbaek et al.
1998] and that Ob-Ra on the choroid plexus clears leptin from the CSF [Tartaglia et al.
1995]. Leptin can be internalized by a coated-pit mechanism by binding to Ob-Ra and
partitioned into a lysosomal compartment where it can then be degraded [Uotani et al.
1999]. In addition, Ob-Ra is the receptor in the kidney thought to play a role in clearance
of leptin from the circulation [Cumin et al. 1996], possibly by glomerular filtration.
Ob-Rb is highly expressed in the hypothalamus [Lee et al. 1996] and is the
isoform of the receptor through which leptin exerts a biological effect [Tartaglia et al.
1995]. The hypothalamus is the only tissue in which the long form of the leptin receptor
22
is more abundantly expressed than the short form [Tartaglia 1997]. Within the
hypothalamus, Ob-Rb is strongly immunoreactive in the arcuate, paraventricular,
surpraoptic [Matsuda et al. 1999], ventromedial, and dorsomedial nuclei [Mercer et al.
1996]. Ob-Rb is also found in brain, cerebellum, testes, adipose tissue [Lee et al. 1996],
and pituitary [Jin et al. 1999; Jin et al. 2000]. In pituitary, leptin receptor is found in 70%
of ACTH cells, 21% of GH cells, 33% of FSH cells, 29% of LH cells, 32% of TSH cells,
and 3% of prolactin cells [Jin et al. 1999].
Recently, a soluble isoform of the leptin receptor (Ob-Re) has been characterized
which has an affinity similar to that of the membrane-bound Ob-Rb [Liu et al. 1997]. It
has been proposed that the role of Ob-Re is to regulate leptin bioactivity by protection
from degradation, slow the clearance process, and inhibit binding to Ob-Rb [Liu et al.
1997]. This soluble receptor is potentially the same protein as a plasma binding molecule
[Liu et al. 1997] and has been shown to bind the majority of circulating leptin in mice
and humans [Houseknecht et al. 1996].
The other short forms of the leptin receptor are far less characterized. Ob-Rc and
Ob-Rd are expressed in heart, testes, adipose tissue, and spleen [Lee et al. 1996]. Ob-Rf
is localized in the brain, liver, stomach, kidney, lung, heart, thymus, spleen, and
hypothalamus [Wang et al. 1996]. More work needs to be done to elucidate the roles
these receptors play in leptin bioactivity.
23
Table 1-3: Leptin Receptors
Receptor Length Localization
Ob-Ra
Short
Choroid plexus, leptomeninges,
brain microvessels, brain,
hypothalamus, testes, adipose tissue,
liver, stomach, kidney, heart, lung
Ob-Rb
Long
Ob-Rc and
Ob-Rd
Ob-Re
Ob-Rf
Short
Short
Short
Hypothalamus, brain, cerebellum,
pituitary, testes, adipose tissue
Heart, testes, adipose tissue, spleen
Soluble - circulates in blood
Brain, hypothalamus, liver, stomach,
heart, kidney, lung, thymus, spleen
Roles
Transports leptin across
blood brain barrier,
degrades and/or clears leptin
from cerebrospinal fluid,
clears leptin from
circulation
Biological effects
?
Regulates leptin bioactivity
?
Leptin Signaling
Members of the class I cytokine receptor superfamily, to which the leptin receptor
belongs, individually demonstrate no signaling capacity, but rather require complexation
with protein kinases to initiate a signal transduction cascade [Tartaglia et al. 1995]. All
membrane-bound isoforms of the leptin receptor have a Box 1 domain [Murakami et al.
1991] and the long form also has a Box 2 domain [Lee et al. 1996; Bjorbaek et al. 1997].
Both Boxes 1 and 2 are required for JAK (Janus kinase or just another kinase) binding
and activation of signal transduction via the signal transducers and activators of
transcription (STAT) pathway [Murakami et al. 1991]. It was initially suggested that the
short forms of the leptin receptor were incapable of signal transduction [Tartaglia et al.
1995].
As stated previously, cytokine receptors cannot initiate a signal transduction
cascade independently [Tartaglia et al. 1995]. Many cytokine receptors are capable of
24
signaling following receptor homooligomerization, and it was predicted that Ob-Rb
followed the same pattern [White et al. 1997]. It was shown that the long form as well as
the short form of the leptin receptor homodimerizes spontaneously [White and Tartaglia
1999] and can form heterodimers upon ligand binding.
The JAK-STAT pathway is common in cytokine signaling [Darnell 1997;
Pellegrini and Dusanter-Fourt 1997]. JAKs are constitutively associated with the
cytoplasmic domain of cytokine receptors. When the appropriate ligand binds to its
receptor, the receptor dimerizes and JAKs are able to phosphorylate each other. Upon
this activation, JAKs phosphorylate the receptor and various intracellular transcription
factors. One such factor is STAT. Src homology 2 (SH2) domains, found on all STATs,
promote binding of STATs to phosphorylated receptors [Darnell et al. 1994; Ihle and
Kerr 1995; Schindler and Darnell 1995], which subsequently results in phosphorylation
of STATs by JAKs. Phosphorylated STATs dimerize, translocate to the nucleus, and
induce gene transcription to produce a biological effect. The long form of the leptin
receptor has 3-5 sites where tyrosine phosphorylation may occur [Bjorbaek et al. 1997].
JAKs phosphorylate these sites and allow the interaction of STATs with the receptor.
Leptin is also capable of signal transduction through pathways that involve the
mitogen activating protein (MAP) kinase and insulin receptor substrate I (IRS-1)
[Bjorbaek et al. 1997; Murakami et al. 1997; Yamashita et al. 1998]. JAK activation is
required to initiate the MAP kinase transduction cascade [Wang et al. 1995; Winston and
Hunter 1996]. JAK phosphorylates Shc, which then activates Ras, and the cascade is
initiated. Alternatively, JAK mediates IRS-1 phosphorylation [Argetsinger et al. 1995;
Johnston et al. 1995], thereby activating Ras. Recently, however, it was found that the
25
short form of the leptin receptor, using the Box 1 motif, was capable of signal
transduction through a pathway in which Box 2 was not involved [Bjorbaek et al. 1997;
Murakami et al. 1997; Yamashita et al. 1998]. The Box 1 pathway phosphorylates JAK2
and IRS-1 tyrosine residues and activates the MAP kinase cascade, but is incapable of
STAT activation. The long form of the leptin receptor was better able to activate the
STAT pathway than the short form, however, it was clear that the short form had
signaling capabilities [Bjorbaek et al. 1997].
JAK-STAT transcription activity is rapid and transient [Shuai et al. 1992],
indicating that the pathway is negatively regulated. Suppressors of cytokine signaling
(SOCS) are proteins that are encoded by genes that are activated by the STATs
[Yoshimura et al. 1995; Starr et al. 1997]. There are many members of the SOCS family,
including SOCS-1 through SOCS-7 and CIS (cytokine inducible SH2-containing protein)
[Hilton et al. 1998]. There is differential expression of SOCS in various tissues [TolletEgnell et al.1999] and each member acts in a different way to negatively feedback on
JAK or STAT to limit the intensity and/or duration of activation [Nicholson and Hilton
1998; Starr and Hilton 1998]. Leptin treatment has been shown to induce SOCS-3 and
CIS mRNA in many target tissues [Emilsson et al. 1999]. Excess leptin production, such
as that in obesity, may result in overproduction of cytokine suppression proteins, and as
such is a potential mechanism of leptin resistance [Emilsson et al. 1999].
Leptin Resistance
As a circulating factor informing the brain of the body’s energy stores, leptin
plays a role in the maintenance of body weight homeostasis. Mice with mutations of the
ob gene have reduced levels of circulating leptin and exhibit hyperphagia and obesity.
26
When these animals are given exogenous leptin, their body weight is significantly
reduced. Mice with mutations of the db gene have elevated levels of circulating leptin
but, like the ob mutants, experience hyperphagia and obesity. Leptin treatment in the db
mutants is ineffective. Human obesity is not often the result of a mutation, as far as is
currently known, so comparisons with these animal models is not entirely accurate;
however most obese humans exhibit dramatically elevated levels of circulating leptin
[Zhang et al. 1994; Maffei et al. 1995b] and ob mRNA [Considine et al. 1996b]. In this
respect, humans more closely resemble the db/db mouse than the ob/ob mouse and, in
general, are insensitive to the effects of leptin [Frederich et al. 1995a; Maffei et al.
1995b; Halaas et al. 1997]. The problem in humans is not the production of leptin, but
the response to leptin.
When the brain is unable to respond to elevated leptin levels, it is referred to as
resistance [Flier and Elmquist 1997]. It has been suggested that this so-called leptin
resistance is a major cause of obesity [Frederich et al. 1995a; Frederich et al. 1995b;
Maffei et al. 1995b; Considine et al. 1996b]. There are many potential mechanisms of
leptin resistance. Transport of leptin across the blood-brain barrier is unidirectional from
blood to brain [Banks et al. 1996] and occurs by a saturable process [Caro et al. 1996a].
The saturable transport of leptin into the brain may be a critical component in resistance
[Tartaglia 1997]. In a recent study in mice, it was shown that central administration of
leptin to peripherally resistant animals resulted in significant reductions in food intake
and body weight [Van Heek et al. 1997], suggesting that leptin may lose its ability to
cross the blood-brain barrier in the resistant state. In addition, it was shown that the
cerebrospinal fluid (CSF)-to-serum leptin ratio is lower in obese humans compared to
27
lean [Caro et al. 1996a; Schwartz et al. 1996b]. Other investigators have also
demonstrated an inability of leptin to cross the blood-brain barrier in resistant animals
[Banks et al. 1996; Caro et al. 1996a; Schwartz et al. 1996b; Van Heek et al. 1997].
However, all aspects of leptin resistance cannot be explained by its inability to cross the
blood-brain barrier. The Koletsky rat, a leptin receptor null model, has leptin CSF levels
similar to controls [Wu-Peng et al. 1997], indicating that leptin can enter the brain
independently of leptin receptors and other transport mechanisms [Bjorbaek et al. 1998].
Another mechanism of leptin resistance may be explained by the inability of the
hypothalamus to detect leptin [Tartaglia 1997] due to defective hypothalamic leptin
receptors [Considine et al. 1996a ; Dawson et al. 1997]. Considine et al. report that the
full-length leptin receptor is expressed in the human hypothalamus [Considine et al.
1996a]; perhaps leptin receptors are functional but downregulated in response to
hyperleptinemia. The possibility that there is a blocked leptin receptor has also been
suggested [McGregor et al. 1996]. Alternatively, leptin resistance may occur
downstream of the receptor in the signal transduction pathway [Tartaglia 1997].
There are varying stages of leptin resistance among different strains of obese mice
[Halaas et al. 1997]. In a study comparing many strains of obese mice, leptin given
peripherally was effective in lean mice and in one strain of obese mice but at a much
higher dose. In addition, leptin given directly into the brain was effective in two of three
strains of obese mice tested but at dramatically different doses. The significance of this
study is that there are likely to be variations of leptin resistance in obese humans as well.
28
Human Leptin Mutations
Although the majority of humans experience obesity due to leptin resistance
[Frederich et al. 1995a; Frederich et al. 1995b; Maffei et al. 1995b; Considine et al.
1996b], there are some instances of mutations of the leptin or leptin receptor. Two
cousins in a Pakistani family, normal weight at birth, developed severe early-onset
obesity [Montague et al. 1997]. No one in the families of either of the children was
obese. It was found that the obesity is due to a deletion mutation of a single nucleotide of
the leptin region of the obese gene, which results in a premature stop codon. Both
children are homozygous for the mutation; their parents are heterozygous. This mutation
caused the creation of a form of leptin that cannot be secreted; serum leptin levels are
nearly undetectable. In addition, the low levels of serum leptin present lack the disulfide
bond that produces bioactivity [Rau et al. 1999]. The children are hyperphagic but
display no noticeable impairment of basal energy expenditure [Montague et al. 1997].
Mutations of the obese gene, as have been described here, are rare [Reed et al. 1996]. It
has been shown, however, that mutations on chromosome 6 in the region of the obese
gene predispose toward extreme obesity [Clement et al. 1996].
In a Turkish family, congenital leptin deficiency resulted in severely obese
individuals. This deficiency is the result of a missense mutation in the leptin gene
[Strobel et al. 1998], which results in impaired secretion of leptin. The family members
homozygous for the mutation are severely obese, hyperphagic, and have low sympathetic
tone. The mutations in both the Pakistani and Turkish families are identical to that in the
ob/ob mouse.
A Kabilian family with severely obese individuals are homozygous for a nonsense
mutation in the leptin receptor [Clement et al. 1998]; this mutation being homologous to
29
that in the db/db mouse and fa/fa rat. The mutation results in a protein that lacks the
transmembrane and intracellular domains of leptin receptor. These subjects develop
obesity within the first year of life, are hyperphagic, and have high circulating levels of
leptin. As in the Pakistani and Turkish families, the members of the Kabilian family who
are heterozygous for the mutation do not develop the phenotype.
Leptin Treatment in Humans and Leptin Gene Therapy
As would be predicted from studies in which leptin was administered to ob/ob
mice, leptin treatment has been shown to be effective in an individual with congenital
leptin deficiency. The nine-year old female cousin from the Pakistani family was treated
with recombinant leptin for 12 months [Farooqi et al. 1999]. The treatment resulted in
weight reduction due to loss of fat and negative energy balance due to reduced food
intake. After 2 months of therapy, leptin antibodies were detectable in the plasma; they
did not, however, appear to interfere with the response to the treatment.
Leptin treatment may also be effective in patients whose obesity is not the result
of genetics. In a clinical trial in which leptin was given to both obese and lean subjects
[Heymsfield et al. 1999], leptin was effective at producing weight loss in a dosedependent manner over a 4-week period. In addition, leptin was effective for 24 weeks in
obese subjects. Weight loss was primarily due to fat loss [Heymsfield et al. 1999], as
was seen in the Pakistani female [Farooqi et al. 1999]. The results of this study are
promising because they indicate that, in instances in which resistance is not the result of a
mutation, high doses of exogenously administered leptin may overcome leptin resistance
and help obese individuals lose weight [Heymsfield et al. 1999].
30
It has been shown that leptin gene therapy is effective in animals [Chen et al.
1996; Murphy et al. 1997; Shimabukuro et al. 1997]. In animals that were made
hyperleptinemic with an adeno-associated virus (AAV), triglyceride content in several
tissues which express leptin receptor was reduced [Shimabukuro et al. 1997]. Murphy et
al. [1997] created a recombinant AAV, which expresses leptin both in vivo and in vitro.
A single intramuscular injection of this rAAV to ob/ob mice resulted in correction of
metabolic defects, obesity, diabetes, elevated food intake and body weight,
hyperinsulinemia, and insulin resistance for as long as six months [Murphy et al. 1997].
Chen et al. [1996] also induced chronic hyperleptinemia in normal adult male rats by the
aid of adenovirus-mediated gene delivery. In these animals, there was a reduction in food
intake, no increase in body weight, and disappearance of adipose tissue. These results,
taken together with results from leptin treatment in humans, suggest that leptin gene
therapy may be beneficial in the treatment of human obesity.
Growth Hormone
Growth hormone (GH) is a 21.5 kilodalton, 191 amino-acid polypeptide that is
secreted episodically from the somatotropic cells of the anterior pituitary gland [Martin et
al. 1978]. The secretion of GH is under direct hypothalamic control mediated by two
neuropeptides: growth hormone releasing hormone (GHRH) and somatotropin-release
inhibiting hormone (SRIH) [Martin and Millard 1986; Millard 1989]. GHRH,
synthesized and released from neurons located in the hypothalamic arcuate nucleus, is
responsible for the high amplitude GH secretory episodes in both man and rats. SRIH, on
the other hand, mediates the prolonged intervals whereby very low levels of GH secretion
occur. SRIH neurons are confined to the periventricular nucleus of the hypothalamus
31
[Kiyama and Emson 1990] in both man and rodents. GHRH and SRIH are secreted
consistently into hypophyseal portal blood, and each also display a periodic surge every
3-4 hours. Each neuropeptide is secreted 180° out of phase with the other [Tannenbaum
and Ling 1984]. SRIH regulates GHRH secretion [Tannenbaum 1994] and the release of
GHRH and SRIH are, in turn, regulated by a host of other neuropeptides and putative
neurotransmitters, including neuropeptide Y (NPY). In addition, synthetic growth
hormone releasing peptides (GHRPs) have been shown to be more potent at stimulating
GH release than GHRH [Bowers et al. 1991; Bowers 1994]. GHRPs synergize with
GHRH to release GH, which suggests that GHRH and GHRPs act at different pituitary
somatotrope receptors and through different second messenger pathways. GHRPs also
regulate GH by antagonizing the actions of SRIH [Ghigo et al. 1997].
In addition to regulation by GHRH, SRIH, and GHRP, GH secretion is altered in
response to many other neuropeptides, neurotransmitters, hormones, and physiological
conditions (Tables 1-4 and 1-5). These influences may occur directly at the level of the
pituitary or by control of the hypothalamic regulators of GH. When considered to its full
extent, the regulation of GH secretion is very complex. To further complicate the matter,
the regulation of GH is not always similar among species or within various disease states.
For example, the regulation of GH by histamine [Netti et al. 1981; Knigge et al. 1990],
bombesin [Pontiroli and Scarpignato 1986; Scarpignato et al. 1986; Benitez et al. 1990],
neuromedin C [Houben and Denef 1991], excitatory amino acids [Mason et al. 1983],
starvation [Shibasaki et al. 1985], and exercise [Felsing et al. 1992; Butkus et al. 1995]
have been demonstrated only in rats or have differential effects in man and rodents. In
addition, regulation of GH by thyrotropin releasing hormone [Czernichow et al. 1976;
32
Mueller et al. 1977; Valentini et al. 1989; Giustina et al. 1995] and dopamine [Liuzzi et
al. 1974; Chihara et al. 1979; Peillon et al. 1979; Kitajima et al. 1986; Schober et al.
1989] is altered in different disease states.
Table 1-4: Mediators that Inhibit GH Secretion in Both Man and Rodents
Mediator
Type
Reference
SRIH
Neurohormone
Arginine vasopressin
Neurohormone
Martin and Millard 1986;
Millard 1989
Martin et al. 1978a
β2 adrenergic agonists
Neurotransmitter
Mauras et al. 1987
Nicotinic muscarinic
agonists
Plasma fatty acid
concentration
Age
Neurotransmitter
Mendelson et al. 1981
Physiological
condition
Physiological
condition
Physiological
condition
Imaki et al. 1985
Obesity
Iranmanesh et al. 1994;
Veldhuis and Iranmanesh 1996
Iranmanesh et al. 1994;
Veldhuis and Iranmanesh 1996
Table 1-5: Mediators that Stimulate GH Secretion in Both Man and Rodents
Mediator
Type
Reference
GHRH
Neurohormone
Martin and Millard 1986;
Millard 1989
GHRP
Synthetic
Bowers et al. 1980;
oligonucleotides
Bowers et al. 1984
Galanin
Neuropeptide
Murakami et al. 1989;
Giustina et al. 1992
Opiates
Neuropeptides
Delitala et al. 1983;
Murakami et al. 1985
Neurotransmitter
Lancranjan and Marback 1977;
α2 adrenergic agonists
Miki et al. 1984
Cholinergic muscarinic
Neurotransmitter
Locatelli et al. 1986
agonists
Serotonin
Neurotransmitter
Imura et al. 1973;
Murakami et al. 1986
GABA
Neurotransmitter
Cavagnini et al. 1977;
Acs et al. 1987
Testosterone
Hormone
Veldhuis et al. 1995a;
Mauras et al. 1996
33
GH pulses can be detected in the blood every 3-5 hours in humans, baboons
[Steiner et al. 1978], monkeys [Quabbe et al. 1981], rabbits, dogs, and rats [Martin
1978]. There are binding proteins for GH found in the plasma. These proteins, GHBPs,
correspond to the extracellular domain of the GH receptor [Leung et al. 1987] and, as
such, bind GH with high affinity. The role of these circulating binding proteins is to
protect GH from premature degradation or clearance [Baumann et al. 1987] and thus
increase its half-life, which is 17-45 minutes [Martin et al. 1978].
GH secretion is associated with the onset of sleep and slow wave sleep [Goldstein
et al. 1983] and levels fluctuate throughout the life span. GH levels are very high in
neonates due to elevated daytime and nighttime bursts of GH [de Zegher et al. 1993].
Values fall shortly after birth and remain stable from day to day in prepubertal children
[Martha et al. 1996]. During puberty, pulsatile GH secretion is amplified as much as 3fold [Mauras et al. 1996]. Following puberty, young adults experience a fall in GH to
levels equal to or lower than those in the prepubertal stage [Martha et al. 1996]. These
levels slowly fall throughout adulthood and are greatly reduced in aged individuals
[Veldhuis and Iranmanesh 1996].
GH has many important roles in the body. It is mitogenic, increasing RNA and
DNA synthesis and inducing cell division [Merimee 1979]. It is anabolic, incorporating
amino acids into proteins, increasing muscle size and strength, and lengthening and
widening bones [Merimee 1979]. GH also regulates body composition through nitrogen
sparing and lipolysis [Ho et al. 1996] directly at the level of the adipocyte [Fagin et al.
1980; Vikman et al. 1991]. In obese animals, GH burst amplitudes are reduced [Veldhuis
et al. 1991] and the resulting (decreased) levels of GH may not be sufficient to affect
34
lipolysis. Ahmad et al. found that in genetically obese Zucker (fa/fa) rats, GH and
GHRH and their messages were decreased by the age of five weeks [Ahmad et al. 1993].
When the rats were given recombinant human GH (rhGH), a decrease in body weight
was observed.
There are various signaling pathways by which GH works, but the main pathways
are JAK-STAT [Argetsinger and Carter-Su 1996] and SOCS [Tollet-Egnell et al.1999].
Once secreted into the plasma, GH stimulates the release of insulin-like growth factor I
(IGF-1). IGF-1 is released in a continuous manner and circulates in the plasma bound
predominantly to one of its binding proteins, IGF binding protein-3 (IGFBP3) [Lamson et
al. 1991]. Both IGF-1 and its binding protein have relatively long half-lives in this
complex [Mohan and Baylink 1996]. Since GH stimulates IGF-1 and because of its
continuous release and relatively long half-life, IGF-1 is considered a good indicator of
GH status over time [Blum et al. 1990]. IGF-1 does not appear to have circadian
rhythmicity, so time of sampling is not important [Sara and Hall 1990]. The main source
of circulating IGF-1 is the liver [Sara and Hall 1990]. Regulation of liver IGF-1 is
strongly regulated by GH, but fasting and refeeding also have IGF-1 regulatory roles
[Philipps et al. 1989]. IGF-1 is also produced in many other tissues of the body in which
it acts as a paracrine factor [Hall and Bozovic 1969]. GH is a primary regulator of IGF-1
in extrahepatic tissues such as heart, lung, and pancreas [Roberts et al. 1987]. Other IGF1 regulators in extrahepatic tissues include prolactin [Murphy et al. 1988] and various
growth factors and trophic hormones [Clemmons and Shaw 1983; Clemmons 1985; Sara
and Hall 1990]. IGF-1 is also produced locally in response to neural [Hansson et al.
1986], arterial [Hansson et al. 1987], and skeletal muscle injury [Jennische et al. 1987].
35
GH has both direct effects in the body and indirect effects mediated by IGF-1.
Some of the direct effects of GH include lipolysis [Fain et al. 1965] and gluconeogenesis
[Dawson and Hales 1969]. Some of the effects mediated by IGF-1 include many of the
growth-promoting effects in muscle, cartilage, and bone, such as DNA synthesis, cell
proliferation, and protein synthesis [Schoenle et al. 1982]. Green et al. [1985] proposed a
dual model of GH action which suggested that GH stimulates differentiation of precursor
cells and IGF-1 then stimulates growth.
Excess of Deficiency of Growth Hormone
Excess and deficient GH secretion result in pathological conditions that can be
corrected pharmacologically. For example, GH hypersecretion usually occurs as the
result of a pituitary adenoma [Hansen et al. 1994]. When this hypersecretion begins
before puberty, gigantism occurs. When the hypersecretion starts after puberty, the result
is acromegaly. In both conditions, individuals experience elevated IGF-1, insulin
resistance, and impaired glucose tolerance [Quabbe and Plockinger 1996]. There is also
increased fat mobilization [Weil 1965], amino acid retention, and stimulated protein
synthesis [Russell-Jones et al. 1993], which result in decreased fat mass and increased
lean mass [Salomon et al. 1993]. Muscle strength is not necessarily greater. Increased
total body water and extracellular fluid result in mild arterial hypertension [Quabbe and
Plockinger 1996]. Bone turnover is increased [Lieberman et al. 1992]. In gigantism,
bones are widened as well as lengthened. In acromegaly, the GH excess occurs after
puberty when the epiphyseal plates fuse and bones can no longer grow in length. A
thickening of bones occurs in acromegaly. In addition, there is excess growth of cartilage
and soft tissue cell mass [Quabbe and Plockinger 1996]. Hypersecretion of GH can be
36
treated long-term with an analogue of SRIH, octreotide [Sassolas 1992]. Octreotide
therapy reduces GH and IGF in many patients with acromegaly [Hansen et al. 1994] and
these reductions result in normalization of lean body mass, body fat, extracellular water
content [Bengtsson et al. 1989; Bengtsson et al. 1990], and joint pain [Sassolas 1992].
However, body cell mass remains high [Bengtsson et al. 1990] and there is an increased
incidence of gallstones [Sassolas 1992].
Individuals deficient in GH experience increased truncal fat mass and decreased
lean mass [DeBoer et al. 1992]. There are also symptoms of osteopenia [Holmes et al.
1994], adverse lipid profiles [Cuneo et al. 1993], glucose intolerance and resistance
[Beshyan et al. 1994; Johansson et al. 1995], and reduced exercise capacity [Nass et al.
1995]. Finally, there is a general reduction in the quality of life [Bjork et al. 1989;
McGauley 1989; Rosen et al. 1994]. Growth hormone replacement increases circulating
IGF-1 values. It normalizes lean and fat mass [Binnerts et al. 1992], redistributing
adipose tissue from abdominal to peripheral depots [Bengtsson et al. 1992]. Patients gain
muscle [Bengtsson et al. 1990] through increased protein synthesis. There are also
increases in bone turnover [Binnerts et al. 1992]. In addition, individuals undergoing GH
replacement therapy experience improved quality of life [McGauley et al. 1990] and
cognitive functioning [Almqvist et al. 1986]. Side effects can include transient water
retention [Binnerts et al. 1992] that can be eliminated by lowering the dose of GH.
Growth Hormone and Obesity
It is widely recognized that GH is attenuated in obesity. A number of negative
correlations have been demonstrated between measures of body mass and GH, including
BMI vs. GH release, amplitude, and half-life [Iranmanesh et al. 1991; Veldhuis et al.
37
1995b] and percent body fat vs. GH release and half-life [Veldhuis et al. 1995b]. The
somatotrope secretory capacity is reduced in obesity [Maccario et al. 1997]; obese
individuals experience reductions in frequency, amount, and duration of GH secretion
and shorter plasma GH half-life [Veldhuis et al. 1991]. The exact mechanism(s) of the
reduction of GH in obesity is unknown [Scacchi et al. 1999], however, it is known that
there are multiple players involved.
There may be a hypothalamic contribution to attenuation of GH in obesity. It was
previously suggested that elevated SRIH contributed to attenuated GH in obesity
[Cordido et al. 1989; Tannenbaum et al. 1990]. However, another study showed a
decreased CSF level of SRIH in obese patients [Brunani et al. 1995]. It was also
previously hypothesized that reductions in GHRH caused the GH attenuation in obesity
[Tannenbaum et al. 1990; Ahmad et al. 1993]. This hypothesis was also challenged
when it was demonstrated that GHRH concentrations did not differ between obese and
nonobese subjects [Brunani et al. 1995]. However, when given exogenous GHRH,
overweight subjects exhibit a blunted rise in GH whether GHRH was given by
intravenous bolus [Williams et al. 1984], continuous intravenous infusion [Davies et al.
1985], or pulsatile intravenous administration [Kopelman and Noonan 1986]. In
addition, the response of GH to GHRPs was reduced in obese patients compared to lean
[Cordido et al. 1993]. Obviously, more work needs to be done to fully elucidate the role
of the hypothalamic regulatory peptides on GH in the obese state.
Another potential mechanism of attenuated GH in obese patients is decreased
half-life, which may be explained by the reduction of circulating GHBPs in obese
patients [Hochberg et al. 1992; Argente et al. 1997; Kratzsch et al. 1997]. There is a
38
positive correlation between both BMI [Hochberg et al. 1992] and percent body fat
[Kratzsch et al. 1997] and GHBPs in obesity. GHBPs act to protect GH from
degradation and thus extend its half-life, so it is logical that reductions in the amount of
circulating GHBPs result in shorter GH half-life. In addition, metabolic clearance rate of
GH is increased in obese individuals [Veldhuis and Iranmanesh 1996].
Obese individuals also demonstrate a reduced number of IGF-1 receptors and
reduced binding [Hochberg et al. 1992]. There is some controversy as to the effects of
obesity on IGF-1 levels. There have been studies demonstrating both reduced [Argente et
al. 1997] and elevated [Hochberg et al. 1992] plasma IGF-1 levels in obesity, but
reductions in receptor binding limit the activity of IGF-1 either way.
There are many circulating factors that feedback on GH to reduce its secretion
and which may play a role on the effects of GH in obesity. For example, it is known that
circulating insulin feedsback negatively on GH secretion [Lanzi et al. 1997]. Since
obesity is associated with hyperinsulinemia [Polonsky et al. 1988], it is tempting to
speculate that this negative feedback may play a role in inhibiting GH. However, there
are diseases in which insulin is elevated and GH is normal [Maccario et al. 1996]. In
addition, reduction of hyperinsulinemia in obesity does not normalize GH [Chalew et al.
1992]. Non-esterified fatty acids (NEFA) are another example of circulating factors that
feedback to inhibit GH. Upon lipolysis, NEFA are released into circulation. This
increase in NEFA has been shown to negatively feedback at both the level of the pituitary
[Casanueva et al. 1987] and the level of the hypothalamus to reduce GH secretion.
Obese individuals exhibit elevated plasma NEFA [Opie and Walfish 1963; Golay et al.
1986], adding another potential mechanism for the attenuation of GH in obesity.
39
Promisingly, the attenuation of GH is obesity is lessened with the reduction in
body weight. Weight loss restores both spontaneous and GHRH-stimulated GH release
[Williams et al. 1984] and normalizes elevated GHBP levels [Rasmussen et al. 1996;
Argente et al. 1997]. In addition, GH treatment of obese subjects results in desirable
effects on body mass [Jorgensen et al. 1994; Richelsen et al. 1994].
Leptin and Growth Hormone
It has recently been shown that circulating leptin and GH levels are related.
Studies demonstrated a normalization of elevated leptin levels upon GH replacement in
GH-deficient adults [Florkowski et al. 1996, Fisker et al. 1997], most likely a
consequence of the decrease in body fat which occurred as a result of the lipolytic effects
of GH. In addition to regulation of leptin by GH, there is regulation of GH by leptin.
Leptin induces spontaneous and GHRH-induced GH secretion [Tannenbaum et al. 1998],
at least partially by inhibiting SRIH release. Carro et al. [1997] demonstrated that leptin
antiserum decreased GH amplitude and nadir in rats suggesting that normal levels of
leptin are required for normal GH secretion. They also showed that in fasted animals
with low leptin and low GH, administration of exogenous leptin normalized GH
secretion.
The effect of leptin on GH may occur at the hypothalamic level. Leptin receptors
colocalize with GHRH in neurons [Hakansson et al. 1998] and are also found in the
periventricular nucleus [Mercer et al. 1996] where SRIH neurons are located [Kiyama
and Emson 1990]. When administered to fasted rats, leptin prevented the inhibition of
GHRH mRNA that is normally seen in conjunction with fasting [Dieguez 1998; Carro et
al. 1999]. When incubated with rat hypothalamic neurons, leptin decreased basal SRIH
40
mRNA and secretion [Quintela et al. 1997]. The regulation of either, or both, of these
two hypothalamic GH regulatory factors by leptin may result in indirect regulation of GH
by leptin. A direct regulation of GH by leptin may also occur at the pituitary level. It has
recently been shown that leptin and leptin receptors are produced in the anterior pituitary
of humans [Jin et al. 1999] and rodents [Jin et al. 2000].
One potential mechanism by which leptin may regulate GH is via neuropeptide Y
(NPY). NPY is a 36-amino acid peptide isolated in 1982 by Tatemoto et al. that has
considerable evolutionary conservation [Tatemoto et al. 1982]. NPY is the neuropeptide
most abundantly found in the brain [Chronwall et al. 1985] with high densities in and
complex networks throughout the hypothalamic nuclei, including the arcuate,
paraventricular, and periventricular nuclei [Chronwall et al. 1985; DeQuait and Emson
1986]. NPY neurons often colocalize with hormones and norepinephrine. In the arcuate
nucleus, NPY neurons colocalize with GHRH and in many other brain areas, NPY and
SRIH neurons colocalize [Vincent et al. 1982; De Quiat and Emson 1986; Fuxe et al.
1989; Okada et al. 1993].
NPY plays roles in reproduction [Kalra et al. 1989], circadian rhythmicity, and
cardiovascular control [Tatemoto 1989], but perhaps the stimulation of feeding is the best
know function of NPY. When given centrally, NPY invokes a robust food intake in male
and female rats [Kalra et al. 1989] during normal nighttime feeding and during daylight
[Clark et al. 1985]. The response is observed within 15 minutes, and is dose-dependent.
NPY specifically increases the intake of carbohydrates and is thought to act
physiologically at times of energy depletion [Leibowitz 1989]. NPY levels are increased
in animals that do not have leptin [Wilding et al. 1993] or functional leptin receptors
41
[Stephens et al. 1995]. In a study in which NPY-knockout mice were used it was shown
that leptin remained effective on the suppression of feeding indicating that this action of
letpin is mediated by pathways independent of NPY [Erickson et al. 1996]. In this study
it was also suggested that NPY opposes the appetite-suppressing effects of leptin.
It is known that (1) fasted animals have low leptin [Mizuno et al. 1996], (2) low
leptin levels increase NPY [Schwartz et al. 1996a], and (3) high NPY inhibits GH [Okada
et al. 1993] by increasing SRIH and by decreasing GHRH [McCann et al. 1989; Rettori
et al. 1990]. In addition, NPY given centrally blunts leptin-induction of GH secretion
[Carro et al. 1998]. It was also shown that the blunted GH that is observed in the fasted
state occurs in conjunction with elevated NPY mRNA [Vuagnat et al. 1998]. When
leptin was administered in this study, the NPY mRNA was reduced concomitant with
normalized GH levels [Vuagnat et al. 1998]. It is therefore possible that, in addition to
the other regulation pathways, leptin regulation of GH is mediated through NPY [Carro et
al. 1997] or that leptin and NPY regulate GH secretion in parallel [Carro et al. 1998].
Objectives
The regulation of GH by leptin and the effects on body homeostasis are the main
topic of investigation for my dissertation. Most of the literature on GH and leptin
interactions reviewed here was published after I developed my hypotheses. I
hypothesized that (1) in the lean animal, circulating leptin stimulates the release of GH by
acting directly at the level of the anterior pituitary and/or indirectly at the level of the
hypothalamus, and (2) animals that are hyperleptinemic and therefore leptin resistant lose
this ability to regulate GH. My work has been summed up by Scacchi et al.:
Leptin, by favouring GH secretion, might reinforce its own biological effects,
chiefly directed (as far as we presently know) at regulating the body fat content.
42
On the other side, the coexistence of high leptin and low GH serum levels in
obesity fits in well with the concept of a leptin resistance in this condition. [1999,
p.263]
CHAPTER 2
GENERAL METHODS
Animals
Male Long-Evans rats (the strain from which the fa/fa rat was derived) were obtained
from Harlan-Sprague Dawley (Indianapolis, IN) and housed individually under standard
temperature and lighting conditions (12 hour light:dark cycle). Individual housing was
required for individual food intake measurements. Water was available ad libitum at all
times. All procedures using animals received prior approval by the Institution's Animal
Care and Use Committee (IACUC).
Diets
The two test diets were obtained from PJ Noyes Company, Inc. (Lancaster, NH). The
high-calorie, high-fat diet consisted of 20-23% fat and 3.7 kilocalories/gram (kcal/g).
The high-calorie, low-fat diet consisted of 2-3% fat and 3.7 kcal/g. Normal rat chow
#5001, obtained from Purina Mills, Inc. (Richmond, VA), consisted of 5% fat and 3.0
kcal/g. It should be noted that the low-fat diet had a calorie content equivalent to that of
the high-fat diet, both of which were higher in calories than the control diet.
Feeding, Pair-feeding, and Body Weight Measurements
Feeding
The animals were allowed an acclimation period of 5 days during which they became
familiar with the new diets and daily handling. After acclimation, each of the diets was
weighed (Mettler scale, PM200, Highstown, NJ) to the nearest 0.1 gram and administered
43
44
daily to each animal. Each subsequent day, the amount of diet remaining was recorded,
including food that was dropped into or through the bottom of the cages. Daily intake
was calculated by subtracting the amount of food remaining from the amount of food
initially administered. Average daily intake was calculated per group.
Pair-feeding
After the acclimation period, the high fat and control rats were fed ad libitum daily. The
low-fat rats were pair-fed the amount in grams ingested by the high-fat group the
previous day. In this manner, the diets of the animals consuming the treatment diets
differed only in the percentage of fat content, not in the number of calories.
Body weight
Body weights of the animals in all three groups were recorded daily to the nearest gram.
The animals were weighed individually in a top loading scale (Taconic Farms, YG-700,
Germantown, NY).
Leptin Challenge Test
The animals were food deprived for 24 hours prior to the test. The leptin challenge test
consisted of a 30 mg/kg subcutaneous bolus of murine leptin (Amgen, Inc., Thousand
Oaks, CA) given to half of the animals in each treatment group (leptin challenge). The
other half of the animals in each group received a bolus of PBS for control. Food (Purina
rat chow #5001, Purina Mills Inc., Richmond, VA) was returned to the animals one hour
post-injection, allowing the animals time to recover following the injection. Food intake
was then measured at 4 hours and again at 24 hours.
45
Alzet Osmotic Minipumps
Alzet osmotic minipumps were used for the continuous administration of murine leptin
(Amgen, Inc., Thousand Oaks, CA) or vehicle over a 2-week (Model 2ML2, Alza
Scientific Products, Palo Alto, CA) or 4-week (Model 2ML4) period. Pump contents
were delivered at a rate of 5 µg/hour or 2.5 µg/hour, respectively. Leptin or PBS
(vehicle) was loaded into each pump under sterile conditions. Pumps were implanted
subcutaneously under methoxyflurane anesthesia. Leptin was stable for the entire 4 week
treatment period. For doses of leptin used, please refer to Chapters 5 and 6.
Right Atrial Cannulation
Anesthesia
Sodium phenobarbital (65 mg/mL, 45-50 mg/kg, Veterinary Laboratories, Inc., Lenexa,
KS) was administered intraperitoneally at a dose of 45-50 mg/kg to produce analgesia,
anesthesia, and muscle relaxation. Atropine sulfate (0.1 mL of 1 mg/mL concentration,
American Reagent Laboratories, Inc., Shirley, NY) was given intramuscularly in order to
prevent the accumulation of fluid in the lungs. If the animals were incompletely
anesthetized, methoxyflurane (Pitman-Moore, Inc., Mundelein, IL) was intermittently
administered via a nose cone.
Preparation
All surgical equipment and cannula hardware were autoclaved or gas sterilized, as
appropriate. The cranial (from between the ears to between the eyes) and neck areas
(unilaterally from slightly caudal to the breastbone to the jaw) of the rats were shaved and
disinfected with betadine and alcohol and the eyes were protected with lubricant.
46
Surgery
In each rat, the right jugular vein was exposed and ligated to stop blood flow. A small
cut was made in the jugular vein and the cannula was inserted and threaded toward the
heart. Pulsation of the cannula indicated that it reached the heart. The cannula was then
pulled back gently just until the pulsation ceased. At this location, the cannula was
secured into position. The cannula was threaded subcutaneously and externalized at the
base of the skull where it was secured with dental acrylic. Also fixed into the dental
acrylic was a snap fastener to later attach to a snap when collecting blood. The cannula
was filled with heparin to prevent coagulation. The neck incisions of the animals were
closed with wound clips.
Recovery
Post-operatively, the animals were placed on heating pads in cages and were closely
monitored until recovery from anesthesia. The animals were returned to their home cages
until blood sampling. The wound clips were removed within 7-10 days of the surgery.
Blood sampling cages
The animals were each placed into the blood sampling cages two days prior to sampling
to allow for acclimation to new surroundings. The cages are large wooden boxes into
which wire mesh cages complete with bedding, food, and water were placed. Tubing was
inserted into the top of the wooden cage, though a protective wire mesh, and attached to
the cannula implanted into the rat, secured with the snap. Through this tubing, blood was
collected without disturbing the rat, thus reducing confounding by stress hormones. Each
cage has a light timer set to the same schedule as the standard housing facility and an
exhaust system for the continual circulation of fresh air. This cage design was approved
by the institution’s IACUC for temporary housing of rodents.
47
Blood Sampling and Tissue Collection
Blood Collection from Cannulae
Blood sampling occurred every 15 minutes for 6 hours from rats in the specialized cages
described above. Before collection of each sample, 400 µL (the calculated amount of
dead space, which includes heparinized saline used to keep the cannula patent) was
withdrawn and saved. A sample of 300 µL was collected and dead space was returned
immediately following the collection of the first sample. The blood was immediately
centrifuged and the plasma was stored at -35°C until use in hormone and other assays.
Red blood cells were resuspended in sterile heparinized saline and returned to the
appropriate animals following collection of the subsequent blood sample and prior to the
return of the dead space. This was done in order to prevent hypovolemia due to
excessive sampling. This method was used to collect all remaining samples.
Blood Collection from Tail Vein
A scalpel was used to remove the tip of the tail from each unanesthetized rat and 1 mL of
blood was collected into a tube. The tail abrasion required no treatment to stop the
bleeding after the collection of blood. The blood was centrifuged at 3200 rpm for 30
minutes (Beckman Centrifuge Model J-6B, Fullerton, CA) and serum was frozen at -35ºC
until use in hormone and other assays.
Blood Collection via Cardiac Puncture
Rats were anesthetized with methoxyflurane anesthesia and a 23-gauge needle was
inserted into the heart to collect 1 mL of blood. The blood was centrifuged at 3200 rpm
for 30 minutes (Beckman Centrifuge Model J-6B, Fullerton, CA) and serum was frozen
at -35ºC until use in hormone and other assays.
48
Trunk Blood Collection
The animals were sacrificed by decapitation and trunk blood was collected by holding the
decapitated rat over a funnel and allowing blood to drain into a large glass test tube.
Trunk blood was centrifuged at 3200 rpm for 30 minutes (Beckman Centrifuge Model J6B, Fullerton, CA) and serum was stored in the -35ºC freezer until use in hormone and
other assays.
Collection of Hypothalamus
The animals were sacrificed by decapitation and the brain was dissected from the skull
using sterile instruments and placed on ice. The hypothalamus was rapidly dissected
away from the rest of the brain using a sharp razor. The hypothalamus was then rinsed,
blotted dry, and weighed before being placed into a sterile tube and snap frozen in liquid
nitrogen. The samples were frozen at -90ºC until reverse transcription and polymerase
chain reaction (RT-PCR) and Western immunoblotting were initiated for leptin receptor.
Pituitary and Organ Weights
In the skull, the clear membrane over the sella turcica was broken with forceps. The
posterior and intermediate lobes of the pituitary were removed and discarded. The
anterior lobe of the pituitary was removed, rinsed, blotted dry, and weighed. From the
body, the liver, testes, kidneys, heart, and adrenals were removed, rinsed, blotted dry, and
weighed.
Cell Culture
GH1 Cells
GH1 cells (American Type Culture Collection, Rockville, MD) are rat pituitary tumor
cells that hypersecrete GH. Cells were grown to 70% confluency in F-12K media
(ATCC, Rockville, MD) supplemented with 1% non-essential amino acids, 1% L-
49
glutamine, 1% nystatin (Gibco BRL, Life Technologies, Grand Island, NY), and either
10% horse serum and 2.5% fetal bovine serum (FBS, Gibco BRL, Life Technologies,
Grand Island, NY) or 12.5% charcoal-stripped FBS (Hyclone, Logan, UT). Cells were
incubated at 37ºC with 5% CO2. Media was changed 2-3 times weekly. GH1 cells were
back-cultured weekly using standard trypsinization procedures to maintain the cell line.
GH1 cells were used in passages 42-44.
Plating Cells
In general, cells were plated at 200,000 cells/well in 24-well plates in a volume of 1
mL/well. Cells were allowed time to adhere, usually 2-3 days, before experimentation.
On the day of each experiment, medium was aspirated from each well and discarded.
Control or leptin-supplemented (murine leptin, Amgen, Inc., Thousand Oaks, CA)
medium was added to appropriate wells. Experiments were completed as described
below.
Five-Day Time -Course
Control or leptin-supplemented (100nM) media was added to appropriate wells on each
plate. Supernatant was collected each day for 5 days and frozen at -35ºC until GH RIA.
Cells were collected and analyzed for DNA content for normalization of GH values and
to control for potentially inconsistent plating densities.
Media Experiment
Cells were plated using media and supplements as described above, but with differences
in serum content. Serum-free media or media supplemented with either 10% horse serum
and 2.5% FBS or with 12.5% charcoal stripped FBS were utilized. The rationale for
these differences in serum supplementation are explored in Chapter 4.
50
Collecting RNA from GH1 Cells
RNA from GH1 cells was collected for RT-PCR amplification of leptin receptor mRNA.
Media was aspirated from the flask and discarded. Trizol (Gibco BRL, Life
Technologies, Grand Island, NY), a reagent designed for the isolation of total RNA from
tissues and cells, was added to the flask (1 mL/10 cm2) and the cells and Trizol were
triturated. Trizol and cells were collected and RNA was extracted as described below.
RT-PCR was performed as described below.
Radioimmunoassays
GH Iodination
125
I-Na (1 mCi, Amersham Life Science, Inc., Arlington Heights, IL) was added to a 10
µL aliquot of 1 mg/mL rGH-I-6 (NIADDK, NIH National Pituitary Agency, Bethesda,
MD) using a lead shielded syringe. Chloramine T (25 µL of 1.5 mg/mL, Sigma
Chemical Co., St. Louis, MO) was added to initiate the reaction and 50 µL sodium
metabisulfite (2.4 mg/mL, Fisher Scientific, Pittsburg, PA) was added 55 seconds later to
terminate the reaction. Bovine serum albumin (100 µL of 100 mg/mL RIA grade, Sigma
Chemical Co., St. Louis, MO) was added to coat the column. The entire solution was
placed on a Sephadex G-75 (Sigma Chemical Co., St. Louis, MO) or Bio-Gel P-60
(BioRad Laboratories, Richmond, CA) column for separation. Each sample was counted
on the Apex Automatic Gamma Counter (ICN Micromedic Systems Model 28023,
Huntsville, AL with RIA AID software, Robert Maciel Associates, Inc., Arlington, MA)
and the fraction with the highest level of radioactivity was saved and diluted for use in
GH radioimmunoassays (RIAs).
51
Growth Hormone RIA
GH was measured in duplicate (cell culture media) or triplicate (plasma) using materials
supplied by Dr. A. F. Parlow and the National Hormone and Pituitary Program
(NIADDK, Baltimore, MD). Values were expressed in ng/mL in terms of the NIADDK
reference preparation rat GH-RP-2. Plasma collected from hypophysectomized rats was
added to the standard curve in the rat plasma assays to correct for non-specific binding.
Hypophysectomized rats have had their anterior pituitaries removed. The plasma,
therefore, lacks GH but has the GH binding proteins. Monkey-anti-rat GH primary
antibody, diluted 1:70,000, and labeled GH, diluted to approximately 12,000 counts/100
µL, were added to the assay and incubated at room temperature for 3 days. On day 4,
goat-anti-monkey secondary antibody, diluted 1:30, was added. Normal monkey serum
(1:200) was added with the secondary antibody to reduce non-specific binding. The
assay was incubated at room temperature for 1 day. On day 5, all tubes except total
counts were centrifuged for 30 minutes (3200 rpm, Beckman Centrifuge Model J-6B,
Fullerton, CA) at 4°C. Supernatant was removed with a vacuum aspirator and pellets
were counted on the Apex Automatic Gamma Counter (ICN Micromedic Systems Model
28023, Huntsville, AL with RIA AID software, Robert Maciel Associates, Inc.,
Arlington, MA) for 1 minute. The GH RIA has an assay sensitivity of 1 ng/mL and a
range of detection from 1 ng/mL to 320 ng/mL.
IGF Iodination
125
I-Na (1 mCi, Amersham Life Science, Inc., Arlington Heights, IL) was added to a 10
µL aliquot 0.25 mg/mL IGF-1 iodination preparation (BACHEM Bioscience, Inc., King
of Prussia, PA) using a lead shielded syringe. Chloramine T (10 µL of 1.0 mg/mL,
Sigma Chemical Co., St. Louis, MO) was added to initiate the reaction. After 45
52
seconds, 200 µL of 100 mg/mL bovine serum albumin (RIA grade, Sigma Chemical Co.,
St. Louis, MO) was added. The entire solution was placed on a Sephadex G-75 column
(Sigma Chemical Co., St. Louis, MO) for separation. Each sample was counted on the
Apex Automatic Gamma Counter (ICN Micromedic Systems Model 28023, Huntsville,
AL with RIA AID software, Robert Maciel Associates, Inc., Arlington, MA) and the
fraction with the highest level of radioactivity was diluted and saved for use in IGF RIAs.
IGF RIA
IGF-1 was extracted from serum by the acid/ethanol procedure. IGF-1 was then
measured by RIA; plasma samples were measured in duplicate for IGF. Values were
expressed in ng/mL in terms of the BACHEM IGF reference preparation (BACHEM
Bioscience, Inc, King of Prussia, PA). Rabbit-anti-rat IGF primary antibody, diluted
1:3,000, and labeled IGF, diluted to approximately 12,000 counts/100 µL, were added to
the assay and incubated at 4°C for 2 days. On day 3, goat-anti-rabbit secondary antibody,
diluted 1:20, was added. Normal rabbit serum (1:50) was added with the secondary
antibody to reduce non-specific binding. The assay was incubated at 4°C for 1 day. On
day 4, all tubes except total counts were centrifuged for 30 minutes (3200 rpm, Beckman
Centrifuge Model J-6B, Fullerton, CA) at 4°C. Supernatant was removed with a vacuum
aspirator and pellets were counted on the Apex Automatic Gamma Counter (ICN
Micromedic Systems Model 28023, Huntsville, AL with RIA AID software, Robert
Maciel Associates, Inc., Arlington, MA) for 1 minute. The IGF RIA has an assay
sensitivity of 0.1 ng/mL and a range of detection of 0.1 ng/mL to 20 ng/mL.
Leptin RIA
Leptin was analyzed with a rat leptin RIA kit (Linco Research, Inc., St. Charles, MO)
which measures both rat and mouse leptin with an assay sensitivity of 0.5 ng/mL and a
53
range of detection from 0.5 ng/mL to 50 ng/mL. Briefly, samples and standards were
aliquoted in duplicate and primary guinea pig antibody, raised against highly purified rat
leptin, was added. The antibody is 100% cross-reactive with rat and mouse leptin and
<2% reactive with human leptin. The tubes were incubated overnight at room
temperature. On day 2, 125I-leptin was added and the tubes were incubated overnight at
room temperature. On day 3, cold precipitating reagent was added and the tubes were
incubated at 4°C for 20 minutes. All tubes except total counts were centrifuged for 40
minutes (3200 rpm, Beckman Centrifuge Model J-6B, Fullerton, CA) at 4°C.
Supernatant was removed with a vacuum aspirator and pellets were counted on the Apex
Automatic Gamma Counter (ICN Micromedic Systems Model 28023, Huntsville, AL
with RIA AID software, Robert Maciel Associates, Inc., Arlington, MA) for 1 minute.
Insulin RIA
Insulin was measured with a rat insulin RIA kit (Linco Research, Inc., St. Charles, MO)
with an assay sensitivity of 0.1 ng/mL and a range of detection from 0.1 ng/mL to 10
ng/mL. Briefly, samples and standards were aliquoted in duplicate and primary guinea
pig antibody, raised highly purified rat insulin, and 125I-insulin were added. The tubes
were incubated overnight at 4°C. On day 2, cold precipitating reagent was added and the
tubes were incubated at 4°C for 20 minutes. All tubes except total counts were
centrifuged for 40 minutes (3200 rpm, Beckman Centrifuge Model J-6B, Fullerton, CA)
at 4°C. Supernatant was removed with a vacuum aspirator and pellets were counted on
the Apex Automatic Gamma Counter (ICN Micromedic Systems Model 28023,
Huntsville, AL with RIA AID software, Robert Maciel Associates, Inc., Arlington, MA)
for 1 minute. The assay is 100% specific for rat, mouse, hamster, human, porcine, and
ovine insulin.
54
Glucose and Triglyceride Assays
Glucose was measured in serum or plasma using a YSI 1500 Sidekick Glucose Analyzer
(Yellow Springs Instrument Company, Inc., Yellow Springs, OH). This instrument
utilizes the glucose oxidase method and the results are linear from 0-800 mg/dL. A 180
mg/dL standard was used. Triglycerides were measured using a kit from Sigma (Sigma
Diagnostics, St. Louis, MO). Triglycerides are hydrolyzed and glycerol is measured with
a qualitative enzymatic method. A 250 mg.dL standard was used and the assay is linear
to a triglyceride level of 1000 mg/dL.
DNA Assay
DNA of GH1 cells was assayed to normalize GH values. After collection of cell culture
media from the 48-well plates (see Cell Culture section), high salt DNA assay buffer
(0.05M sodium phosphate, 2.0M NaCl, 2 mM EDTA, pH 7.4) was added to the cells in
each well and incubated overnight. The samples were collected and sonicated prior to
DNA measurement. The DNA standard curve was derived from calf thymus (Sigma
Chemical Co., St. Louis, MO) and Hoescht florescence emission dye (Molecular Probes,
Eugene, OR) was used. Hoescht dye binds to intact double stranded DNA. Standards
and samples were aliquoted into opaque (white or black) 96-well plates in triplicate.
Hoescht dye (1 µg/mL) was added and the plate was read on a Molecular Devices Type
374 Fluorometer (Labsystems, Finland) using SOFTmax PRO Version 1.3.2 software for
Macintosh at excitation wavelength 320 nm and emission wavelength 520 nm.
55
RNA extraction, RT-PCR, Southern Blot
RNA Extraction
RNA was extracted from hypothalamus and GH1 cells using the phenol-chloroform
method with isopropyl alcohol precipitation. Hypothalami were homogenized and GH1
cells were collected in Trizol reagent (Gibco BRL, Life Technologies, Grand Island,
NY). At the end of the procedure, the pellet was dried and resuspended in RNAse-free
water. Absorbances were read on a spectrophotometer (Model DU-64, Beckman,
Fullerton, CA) to obtain an OD260:OD280 value. OD260:OD280 of 1.7 to 2.1 indicates a
clean RNA extraction.
RT-PCR for Leptin Receptor
RT-PCR for leptin receptor mRNA was optimized using the sense and antisense primers
designed by Zamorano et al. [1997]. The sense and antisense primers were 5’ATGACGCAGTGTACTGCTG and 5’-GTGGCGAGTCAAGTGAACCT, respectively
and amplified a 357-base pair fragment of the long form of the leptin receptor. The
primers were designed using a homologous region in mouse and human leptin receptor
cDNA to amplify the extracellular domain (bp 1274-1630) of the leptin receptor in the rat
(95% homologous with rat). Optimal conditions were identified as 1 mM MgCl 2, 64°C
annealing temperature (Figure 2-1), and 30 cycles (Figure 2-2) following an initial
denaturation phase and followed by a final elongation phase. No plot was made of
density vs. temperature, MgCl 2, or cycle number. The determination of optimal
conditions was made by rough estimation. Negative controls, including a sample lacking
56
the reverse transcription enzyme and a sample lacking RNA, were included (data not
shown).
Figure 2-1: MgCl 2 (mM) and temperature
(°° C) optimization
Lane 1 is the 100 base pair (bp) DNA ladder;
lane 2 is 0.5 mM at 63°C; lane 3 is 1.0 mM at
63°C; lane 4 is 1.5 mM at 63°C; lane 5 is 2.0
mM at 63°C; lane 6 is 0.5 mM at 64°C; and lane
7 is 1.0 mM at 64°C. Based on the results of this
experiment, the conditions used for the
subsequent studies were 1 mM MgCl 2 at 64°C.
Figure 2-2: Cycle optimization
Lane 1 is the mass DNA ladder; lane 2 is
25 cycles; lane 3 is 30 cycles; lane 4 is 35
cycles; and lane 5 is 40 cycles. Based on
the results of this experiment, 30 cycles
were used in subsequent experiments.
Southern Blot and Normalization
The identity of RT-PCR product (Figure 2-3) was confirmed using the internal probe 5’TGCAGCTGAGGTATCACAGG in Southern blot analysis (ECL, Amersham Pharmacia
Biotech, Piscataway, NJ). The inconsistencies in lanes 2 and 3 of the southern blot are
due to relative differences of RNA added to the initial PCR. The amount of leptin
receptor mRNA was normalized with the housekeeping gene cyclophilin. Cyclophilin
did not appear to be affected by the treatment parameters. The cyclophilin sense and
antisense primers used were 5'-GGGAAGGTGAAAGAAGGCAT and 5'GAGAGCAGAGATTACAGGGT, respectively and amplified a 210-base pair fragment
[Zamorano et al. 1997]. Cyclophilin conditions were optimized and found to be the same
as were used for the leptin receptor but half the concentration of RNA was added to the
57
reaction to prevent saturation of cyclophilin. A representative photograph of RT-PCR is
shown in Figure 2-4.
Figure 2-3: Southern blot of leptin receptor
Lane 1 is hypothalamus; lanes 2 and 3 are
GH1 cells; and lanes 4 and 5 are RC cells.
GH1 and RC cells are rat pituitary tumor
cell lines that hypersecrete and
hyposecrete GH, respectively.
Figure 2-4: Representative RT-PCR from
hypothalamus samples
Lane 1 is the mass DNA ladder; lanes 2, 4, 6, and 8
are leptin receptor mRNA; and lanes 3, 5, 7, and 9
are corresponding cyclophilin mRNA.
Protein Extraction and Western Immunoblotting
Protein Extraction
Trizol: Protein was extracted from the same samples from which RNA was extracted
using Trizol reagent (Gibco BRL, Life Technologies, Grand Island, NY). After addition
of phenol/chloroform to isolate RNA, RNA was in the aqueous phase, DNA was in the
interphase, and protein remained in the phenol/chloroform phase. From the
phenol/chloroform phase, protein was precipitated with isopropanol and washed in a
solution containing guanidine hydrochloride. The samples were centrifuged at 7,500 g at
4°C (Beckman Centrifuge Model J2-21, Fullerton, CA) and the pellet was dried and
resuspended in 1% SDS. However, the Trizol reagent does not contain a detergent to
efficiently break cell membranes and release membrane-bound proteins, so additional
samples were extracted in cell lysis buffer (see below) and comparisons were made in
Figures 2-5 and 2-6.
58
Cell Lysis Buffer: Some hypothalami did not require RNA extraction prior to protein
extraction, and therefore were not extracted using the Trizol method. Instead,
hypothalami were homogenized in a phosphate buffer (g tissue x 10 mL buffer), pH 7.4,
containing 5 mM EDTA, 5 mM EGTA, 50 mM NaCl, 5 mM sodium pyrophosphate
decahydrate, 2% Triton X-100, 0.5% SDS (all from Fisher Scientific, Fair Lawn, NJ), 1
mM orthovanadate, 0.1 mM PMSF (both from Sigma Chemical Co., St. Louis, MO), 10
ug/mL leupeptin, 1.0 ug/mL pepstatin, 10 ug/mL aprotinin (all three from Calbiochem,
La Jolla, CA), and 50 mM NaF (Fisher Scientific, Fair Lawn, NJ) using a Brinkman
polytron homogenizer (Brinkman Instruments, Inc., Palo Alto, CA). After
homogenization, samples were kept on ice and vortexed periodically. After 1 hour,
samples were centrifuged (Micro Centrifuge Model 235C, Fisher Scientific, Fair Lawn,
NJ) for 30 minutes and supernatant was frozen at -20°C until further use.
Figure 2-5: Trizol extraction
Lane 1 is the molecular weight marker; lanes
2-6 are hypothalamic samples extracted by the
trizol method. Leptin receptor (long form)
should be near the 126 kDa marker.
Figure 2-6: Cell lysis buffer extraction
Lane 1 is the molecular weight marker; lanes 2-9
are hypothalamic samples extracted by the cell
lysis buffer protocol. Leptin receptor (long form)
is near the 126 kDa marker.
Micro BCA Protein Assay
Micro BCA protein assay is used to measure protein when the samples are suspended in
reagents that interfere with the Coomassie blue of the Bradford protein assay, such as
SDS. A standard curve ranging from 0.063 mg/mL to 1.0 mg/mL was made by serial
dilution in 1% SDS starting with 2 mg/mL albumin standard (Peirce, Rockford, IL). A
59
reference standard (0.0 mg/mL) was also used. Each standard was pipetted into a
colorless 96-well plate in triplicate. Samples were added in duplicate. If dilutions were
required, 1% SDS was used as the diluent. A working reagent was prepared by mixing
reagents MA, MB, and MC (Micro BCA reagents, Pierce, Rockford, IL) at a ratio of
25:24:1. Working reagent was added to standards and samples in each well. The plate
incubated at 37°C for 30 minutes. The plate was then read on the SLT 400 AC Plate
Reader (SLT Lab Instruments, Salzburg, Austria) using dual wavelength (575 nm and
690 nm) with 30 second shake and four parameter standard curve fit. The accompanying
software was DeltaSoft II for Macintosh (BioMetallics, Inc., Princeton, NJ).
Western Immunoblot Analysis for Leptin Receptor
Separation and transfer: The 10 µL protein sample or molecular weight marker
(Kaleidoscope, BioRad Laboratories, Hercules, CA) was combined with 5 µL loading
buffer (0.06 M Tris, 2% SDS, 10% glycerol, 0.025% bromphenol blue, and 5% 2mercaptoethanol), boiled, and loaded onto the pre-cast polyacrylamide-SDS Ready Gel
(BioRad Laboratories, Richmond, CA). For protein loading optimization, see Figure 2-7.
Different volumes of protein were evaluated, 2-9 µL. The best results were seen at a
volume of 9 µL, but the bands were still faint. In all subsequent Westerns, the maximum
amount of protein (10 µL) was loaded. After the Western was completed, µg protein
loaded was calculated. No negative controls were utilized. After loading onto the gel,
the samples were separated by running the gel in a gel electrophoresis apparatus (MiniPROTEAN II Cell, BioRad Laboratories, Hercules, CA) at 125 V for approximately 70
minutes. Samples were transferred to a nitrocellulose membrane (Trans-Blot Transfer
Medium, 0.45 um, 7 x 8.4 cm; BioRad Laboratories, Hercules, CA) at 100 V for 1 hour
using a BioRad Trans Blot Cell apparatus (Hercules, CA).
60
Immunoblotting and detection: The membrane was blocked for 2 hours in 5% nonfat
dry milk in Tris buffered saline with 0.05% Tween (T-TBS) and then incubated with
primary antibody overnight at 4°C. The primary antibody used was rabbit-anti-rat leptin
receptor (long-form) produced to the 18 amino acids near the C-terminus of mouse ObRb (Alpha Diagnostic International, San Antonio, TX). The primary antibody was diluted
1:500 in blocking buffer. After 3 10-minute washes with T-TBS, the membrane was
incubated with the secondary antibody (anti-rabbit IgG-HRP, Santa Cruz Biotechnology,
Santa Cruz, CA) at a dilution of 1:1,000 for 1 hour. Following another washing step, the
blot was exposed to 5 mL of each solution (simultaneously) of SuperSignal
Chemiluminescent Substrate for Western blotting (Pierce, Rockford, IL) for 10 minutes.
The blot was then exposed to radiographic film (Hyperfilm ECL, Amersham Life
Science, Buckinghamshire, England) for 10 minutes. The film was developed in a
Konica Medical Film Processor (QX-70). Optimization for the antibody concentrations
is shown in Figure 2-8. The best combination of primary and secondary antibody shown
in the figure is with each at a dilution of 1:1000, but the resutling bands were faint. A
subsequent test used a more concentrated primary antibody (1:500) with the same
concentration of secondary antibody (1:1000), with better results (data not shown).
These were the conditions used for all subsequent Western analyses.
Analysis: A pooled reference sample was included in each gel to correct for inter-gel
variation. The optical density of the bands was measured using an imaging densitometer
(BioRad Model GS-670, Hercules, CA) and Molecular Analyst Software (version 1.2,
Hercules, CA). The results were normalized according to the amount of protein loaded
(µg) and expressed as a ratio of sample to inter-gel control.
61
Figure 2-7: Protein loading optimization
Lane 1 is the molecular weight marker;
lane 2 is 2 µL; lane 3 is 3 µL; lane 4 is 4
µL; lane 5 is 5 µL; lane 6 is 6 µL; lane 7
is 7 µL; lane 8 is 8 µL; and lane 9 is 9 µL
of a 2.74 µg/µL sample. In subsequent
experiments, 10 µL of each protein
sample was used.
Figure 2-8: Primary antibody concentration
optimization
Lanes 1, 3, 5, and 7 are the molecular weight
markers; lane 2 is 1:1000; lane 4 is 1:1500;
lane 6 is 1:2000; and lane 8 is 1:2500. In
lanes 2, 4, 6, and 8, secondary antibody
concentration was 1:1000. In subsequent
experiments, 1:500 primary antibody and
1:1000 secondary antibody concentrations
were used.
Each leptin receptor protein measured with this Western protocol gave bands of multiple
molecular weights. No pre-immune or pre-absorption samples were run, so the
specificity of the bands is uncertain. The molecular weight of the largest band is what
was expected for the leptin receptor, so it can be assumed to be specific. The lower
molecular weight bands have been seen in studies using different antibodies [Boes et al.
1999], so these are possibly specific as well.
CHAPTER 3
CIRCULATING GROWTH HORMONE LEVELS ARE ELEVATED IN RATS FED A
HIGH-FAT DIET
Introduction
It is well known that the secretion of growth hormone (GH) from the anterior
pituitary gland is episodic. In the past, the rat has been the most widely studied model for
investigation of basic mechanisms that underlie the expression of the GH secretory
pattern in man because of numerous similarities in the GH secretory axes between man
and rats. The regulation of GH secretion is under direct hypothalamic control and is
mediated by the release of a stimulatory neuropeptide, growth hormone releasing
hormone (GHRH) and an inhibitory neuropeptide, somatotropin release inhibiting
hormone (SRIH), also called somatostatin [Martin and Millard 1986; Millard 1989].
In addition to hypothalamic control, metabolic control of GH secretion also shares
similarities in these species. It is clear that circulating GH is dramatically attenuated in
both obese animals [Veldhuis et al. 1991] and obese humans [Williams et al. 1984] and
that this may be directly related to blood-borne factors and their influence on
hypothalamic neurons involved in GH regulation. There are other metabolic controllers
of GH, however, that have the opposite effects between man and rodent species, such as
exercise and fasting.
One possible example of a circulating GH-regulating factor is the obese gene
product, leptin. Leptin is a protein hormone secreted by adipocytes in proportion to body
fat that acts in the brain to help regulate body weight. When there is insufficient leptin or
62
63
impaired leptin recognition by the brain, obesity develops. Circulating leptin influences
and is influenced by metabolic states such as obesity, exercise, and fasting [Mizuno et al.
1996; Ahren et al. 1997]. It is therefore logical to assume that body weight and
consequently leptin may influence GH secretion. An additional argument for this is that
leptin receptors are highly abundant in the arcuate nucleus of the hypothalamus (ARC)
[Mercer et al. 1996], which is also the location of GHRH-producing neurons [Martin
1973].
Leptin, being secreted in proportion to body fat, is higher in the blood of animals
with greater fat deposition. Consequently, it tends to be elevated in the blood of animals
that ingest a diet high in fat [Campfield et al. 1995; Frederich et al. 1995; Masuzaki et al.
1995; Van Heek et al. 1997; Widdowson et al. 1997]. The typical human diet generally
consists of fat in excess of 20% [Wang et al. 1998a], which is much greater that the
laboratory rodent diet that is approximately 5% fat (Purina Mills rodent chow). We
proposed that by placing young rats on a high-fat diet we would elevate circulating leptin
to levels, which would subsequently cause an elevation of circulating GH, assuming that
the length of time the rats were on the diets and the levels to which leptin was raised were
not sufficient to cause obesity/resistance. The results of the present study confirm this
hypothesis.
Methods
Animals
Twenty-five male Long-Evans rats (300-325 g) were obtained from Harlan-Sprague
Dawley and housed individually under standard temperature and lighting conditions. The
animals were divided into three groups each receiving different diets: 9 rats received a
64
high-fat diet (23% fat, 3.7 kcal/g, PJ Noyes Company, Inc., Lancaster, NH), 8 received an
isocaloric, low-fat diet (2% fat, 3.7 kcal/g, PJ Noyes Company, Inc., Lancaster, NH)
which was high in carbohydrates, and 8 controls received normal rat chow (5% fat, 3.0
kcal/g, Purina Mills, Inc., Richmond, VA). The control and high-fat rats were fed ad
libitum. The low-fat rats were pair-fed the amount of food in grams consumed by the
high-fat rats the previous day. Water was available to all groups ad libitum. Body
weight and food intake were measured daily in diet-treated rats and weekly in controls for
the duration of the study.
Cannulation Surgery, Blood Sampling, and Tissue Collection
After four weeks on the various diets, the rats were implanted with right atrial cannulae
under pentobarbital anesthesia (45-50 mg/kg intraperitoneally). The cannulae were
externalized at the base of the skull and secured with dental acrylic. One week after
surgery, 300-400 µL blood samples were collected at 15-minute intervals for a total of 6
hours, beginning between 8 and 9 AM for each rat. Immediately after each 15-minute
sample collection, the blood was centrifuged and the plasma was stored at -35°C until
hormone assays. Red blood cells were resuspended in sterile heparinized saline and
returned to each respective animal following collection of the subsequent blood sample.
This was done in order to prevent hypovolemia due to excessive sampling. After the
completion of sampling, the animals were sacrificed and trunk blood was collected for
analysis of insulin-like growth factor-1 (IGF-1) and leptin. Hypothalami were removed
and frozen at -90°C until reverse transcription and polymerase chain reaction (RT-PCR)
studies were initiated for leptin receptor.
65
Radioimmunoassays
Plasma samples were measured in triplicate for GH using materials supplied by Dr. A. F.
Parlow and the National Hormone and Pituitary Program (NIADDK, Baltimore, MD), as
described [Millard et al. 1981]. Values were expressed in ng/mL in terms of the
NIADDK reference preparation rat GH-RP-6. The GH RIA has an assay sensitivity of 1
ng/mL and a range of detection of 1 ng/mL to 320 ng/mL. Plasma samples were
measured in duplicate for IGF-1. IGF-1 was extracted from trunk blood by the
acid/ethanol procedure and measured by RIA as previously described [Grant et al. 1986].
The IGF RIA has an assay sensitivity of 0.1 ng/mL and a range of detection of 0.1 ng/mL
to 20 ng/mL. Leptin was measured with a RIA kit (Linco Research, Inc., St. Charles,
MO) which measures both rat and mouse leptin with an assay sensitivity of 0.5 ng/mL
and a range of detection from 0.5 ng/mL to 50 ng/mL.
RT-PCR for Leptin Receptor
RT-PCR for leptin receptor mRNA was optimized using the sense and antisense primers
from Zamorano et al. [1997]. The sense and antisense primers were 5’ATGACGCAGTGTACTGCTG and 5’-GTGGCGAGTCAAGTGAACCT, respectively
and amplified a 357-base pair fragment of the long form of the leptin receptor. Optimal
conditions were identified as 1 mM MgCl 2, 64°C annealing temperature, and 30 cycles
following an initial denaturation phase and followed by a final elongation phase. The
identity of RT-PCR product was confirmed using the internal probe 5’TGCAGCTGAGGTATCACAGG in Southern blot analysis (ECL, Amersham Pharmacia
Biotech, Piscataway, NJ). The amount of leptin receptor mRNA was normalized with the
housekeeping gene cyclophilin. The cyclophilin sense and antisense primers used were
5'-GGGAAGGTGAAAGAAGGCAT and 5'-GAGAGCAGAGATTACAGGGT,
66
respectively and amplified a 210-base pair fragment [Zamorano et al. 1997]. The same
conditions were used for cyclophilin as were used for the leptin receptor.
Statistics
Daily and weekly body weight and food intake measurements were compared by twoway repeated measures ANOVA with Student-Newman-Keuls Multiple Comparisons.
Growth hormone profiles were evaluated by Cluster Analysis [Veldhuis and Johnson
1986], which analyzes area under the curve, peak height and width, and valley nadir and
width. Comparisons of IGF-1, leptin, and leptin receptor mRNA were by one-way
ANOVA followed by Tukey Comparisons. Values were significant using p<0.05 unless
otherwise indicated.
Results
Body Weight
Although rats fed the low-fat diet did not lose weight during the study, they did not gain
as much weight as rats fed either the high-fat or control diets (Figure 3-1). There were
significant differences in body weights between low- and high-fat rats beginning on day
10 and remaining for most of the duration of the study and between the low-fat and
control rats on days 10-14 and 18-29. Rats fed the high-fat diet gained significantly more
weight than controls beginning at day 18 and remaining for most of the duration of the
study.
Food Intake
Rats fed the high-fat diet consumed significantly less chow in grams of food than rats fed
the control diet beginning within the first 5 days of the study and remaining for the
duration (Figure 3-2). No comparisons for animals fed the low-fat diet are reported
67
versus either of the other two groups because these rats were pair-fed (in grams of food)
to the high-fat rats. When the data are shown as grams of fat consumed for each
respective diet (Figure 3-2), the high-fat rats consumed significantly more throughout the
study compared to both controls and low-fat rats. In addition, control rats consumed
significantly more fat than low-fat rats for the duration of the study. Control rats
consumed more kcal than rats fed the special diets.
Leptin
At the end of the study, leptin levels were significantly elevated in the rats fed the highfat diet compared to those fed either of the other two diets (Figure 3-3). Although it
appeared that rats fed normal chow had lower leptin values than low-fat rats, the results
were not significant.
Leptin Receptor mRNA
There were no significant differences in leptin receptor mRNA in hypothalamus in
response to any of the different diets (Figure 3-4).
IGF
At the end of the study, IGF-1 levels were significantly elevated in the rats fed the highfat diet compared to those fed either of the other two diets (Figure 3-5).
Growth Hormone Profile
Growth hormone pulses and troughs were analyzed using the Cluster Program [Veldhuis
and Johnson 1986] among the diet-treated and control groups (Table 3-1). It must be
pointed out, however, that each group only averaged two or three values due to sampling
difficulties. Analyses were completed and demonstrated that, overall, the high-fat rats
had the highest total and mean GH under-the-curve values while the low-fat rats had the
lowest (p<0.10). This was most likely due to peak height and valley width. High-fat rats
68
had significantly higher peaks while low-fat rats had significantly lower peaks overall
compared to control (p<0.05). Similarly, the valley nadirs were higher in high-fat rats
and lower in low-fat rats, but these results were not significant. Control rats were in the
middle of the range for both peak height and valley nadir. As expected, because of
elevated pulse amplitudes, the high-fat rats spent less time in the GH trough periods
(valley width, p=0.01) compared to the other groups. Peak width, measured in minutes,
was not significantly different among the groups. Representative profiles for the high-fat
(panel A), control (panel B), and low-fat rats (panel C) are given in Figure 3-6.
Figure 3-1: Body Weight
There were significant differences in body weights between low- and high-fat rats (n=9) beginning on day
10 and remaining for most of the duration of the study and between the low-fat (n=8) and control rats (n=8)
on days 10-14 and 18-29 (2-way repeated measures ANOVA, p<0.05). Rats fed the high-fat diet gained
significantly more weight than controls beginning at day 18 and remaining for most of the duration of the
study.
69
Figure 3-2: Food Intake
When measured in grams (g food), it was shown that rats fed the high-fat diet (n=9) consumed significantly
less chow than rats fed the control diet (n=8) beginning within the first 5 days of the study and remaining
for the duration (2-way repeated measures ANOVA, p<0.05, panel A). No comparisons for animals fed the
low-fat diet (n=8) are reported versus either of the other two groups because these rats were pair-fed in
grams of food to the high-fat rats. When measured in grams of fat consumed (panel B), the high-fat rats
consumed significantly more than the control rats, which in turn consumed significantly more than the lowfat rats (2-way repeated measures ANOVA, p<0.05). When measured in kcal (panel C), normal chow rats
ate more for most of the study, but the values were nearly normalized.
Figure 3-3: Leptin Levels
At the end of the study, leptin levels were significantly elevated (1-way ANOVA, p<0.05) in the rats fed
the high-fat diet (n=8) compared to those fed either of the other two diets (n=6 each).
70
Figure 3-4: Ob-Rb mRNA
There were no significant differences (1-way ANOVA, p<0.05) leptin receptor mRNA in hypothalamus
due to any of the differenct diets (n=6 for each group). Leptin receptor was normalized using the
housekeeping gene cyclophilin.
Figure 3-5: IGF
At the end of the study, IGF-1 levels were significantly elevated (1-way ANOVA, p<0.05) in the rats fed
the high-fat diet (n=8) compared to those fed either of the other two diets (n=6 each).
71
Table 3-1: Growth Hormone Profile of Diet-Treated and Control Rats
Figure 3-6: GH Profile
Representative GH profiles of high-fat (panel A), control (panel B), and low-fat rats (panel C).
72
Discussion
In the mammalian endocrine system, a variety of hormones and their hormonal
and metabolic regulators are intricately intertwined, both centrally and in the periphery.
GH is one such hormone. GH stimulates IGF-1, and separately or in concert, GH and
IGF-1 regulate body composition. In man, GH secretion is enhanced in diabetes [Glass et
al. 1981] and virtually all forms of stress. Both exercise and fasting also increase GH
[Martin and Millard 1986; Borst et al. 1994] while its secretion is attenuated in obesity.
In rats, insulin and corticosterone have a negative effect on circulating GH levels
[Tannenbaum et al. 1981], as do exercise and fasting. Similar to man, GH secretion is
attenuated in obesity in rats [Martin et al. 1983]. The differential regulation of GH
between rodents and man is largely unexplained.
Leptin is a hormone that is regulated by IGF-1 [Bianda et al. 1997] and GH
[Florkowski et al. 1996]. Upon GH replacement in GH-deficient subjects, body fat
decreased resulting in a decrease of elevated leptin [Florkowski et al. 1996]. In another
study, leptin levels were found to be elevated in GH-deficient adults and were normalized
after 1 year of GH therapy [Fisker et al. 1997]. Although the authors claimed there was
no association between the two hormones and that the effects were most likely due to the
decrease in body fat which occurred as a result of the lipolytic effects of GH, there
certainly is evidence that there is at least an indirect relationship. In addition, leptin is
required for maximal blood GH levels [Carro et al. 1997]. Carro et al. [1997]
demonstrated that leptin antiserum decreased GH amplitude and nadir in rats, suggesting
that normal levels of leptin are required for normal GH secretion. They also showed that,
in fasted animals with low leptin and low GH, administration of exogenous leptin
normalized GH secretion.
73
GH has many important roles in the body, including lipolysis. Several
investigators have shown GH receptors on adipocytes [Fagin et al. 1980; Vikman et al.
1991]. In obese animals, GH burst amplitudes are attenuated [Veldhuis et al. 1991].
This reduction of GH in obesity prevents GH-induced lipolysis, perhaps further
contributing to obesity. This is another example of GH-leptin interaction.
In the current study, rats consuming the high-fat diet gained more weight than
controls. The weight gain of the high-fat rats occurred in spite of a food intake in grams
and kilocalories significantly lower than that of controls. These results seem
contradictory but are reasonable when the fat content of each diet is taken into effect.
The grams of fat consumed by the high-fat group exceeded that of the control group,
which in turn exceeded that of the low-fat group. Furthermore, rats that ate the low-fat
diet gained less weight than controls, explained by the differences in overall fat
consumption.
Corresponding to the body weight data, leptin levels were significantly elevated in
the high-fat rats versus the other two groups. This was expected and is probably due to
elevated fat mass in these rats. The levels to which leptin was elevated remained within
the a physiological range, indicating that the animals were not obese and probably were
not leptin resistanct. In addition to elevated leptin, IGF-1 levels were significantly
elevated in the high-fat rats. It has been shown that, in rats, IGF-1 is present in the blood
at relatively constant concentrations dependent on GH secretory status [Donaghue et al.
1990]. IGF-1, therefore, is a good measure of overall GH secretion. Although IGF-1 is
regulated by factors other than GH, it is often used clinically to detect GH secretory
74
problems. The elevation of IGF-1 in response to the high-fat diet may therefore suggest
that GH was also elevated in this group.
A growth hormone profile consisting of samples collected at 15-minute intervals
for 6 hours was established for each of the three groups. Unfortunately, due to sampling
difficulties, the number of animals in each group is insufficient to draw conclusions with
any degree of confidence. However, the results in Table 3-1 strongly suggest that rats fed
the high-fat diet secreted more GH and rats fed the low fat diet secreted less GH when
compared to control rats. Low-fat rats were pair-fed and thus were probably still hungry
when their food supply ran out, however it is unknown how much more food the low-fat
rats would have consumed had they been fed ad libitum. Thus, the implied chronic stress
experienced by these rats potentially due to being in a state of hunger may confound GH
data.
The results of the IGF-1 assays and GH profiles, taken together, suggest that
ingestion of the high-fat diet and the subsequently elevated plasma leptin levels resulted
in increased circulating GH values. Although correlation does not imply causation, it is
possible that the physiological elevations in leptin levels stimulated the release of GH,
either directly or indirectly. The literature suggests that leptin is required for normal GH
release [Carro et al. 1997].
A natural feedback loop involving leptin and GH seems evident. Consider this
greatly simplified physiological situation: when an animal begins to eat more and gain
weight, resulting in hypertrohy and/or hyperplasia of fat cells, more leptin is produced.
This elevation of leptin is sensed by the hypothalamus which decreases feeding and
stimulates metabolism and perhaps directly by the pituitary to enhance GH secretion. GH
75
then acts directly on adipocytes [Fagin et al. 1980; Vikman et al. 1991] to reduce their
size and number, and this with the decreased food intake result in less fat and decreased
or normalized leptin production. When this fall in leptin is sensed by the hypothalamus,
food intake is again increased and GH secretion is reduced in an effort to prevent
lipolysis until normal fat mass has been attained.
In the case presented in the current study, the animals are given a high-fat diet,
and, as expected, gain weight and have elevated circulating leptin. We consequently see
the predicted elevation in GH secretion, mainly observed via IGF-1. The remainder of
the feedback loop described above is not observed, however, in which elevated GH
stimulates lipolysis and decreases leptin secretion. This is probably due to the high fat
content of the high-fat diet, which prevents a loss of body weight and a fall in leptin
levels.
Pathophysiologically, when an animal develops leptin resistance, the ability of the
hypothalamus to detect circulating leptin is attenuated and GH secretion will be
attenuated, as is seen in obesity and which may further contribute to obesity. Assuming
this interpretation is correct, no resistance has yet developed in the current study. First,
the circulating leptin levels, although elevated in rats fed a high-fat diet, remained within
a physiological range. Second, we have seen that GH is still elevated in response to
leptin. Last, we observed no down-regulation of leptin receptor mRNA in the
hypothalamus. In the chapters of this dissertation that study leptin resistance, we the
opposite effects as are demonstrated here: in resistant rats, leptin levels are much higher,
GH is attenuated, and hypothalamic leptin receptor is downregulated.
76
In summary, there is a relationship between leptin and GH in the normal animal.
We placed young rats on a high-fat diet, which resulted in elevated circulating leptin
levels and subsequently elevated endogenous GH and IGF-1 secretion. There are several
mechanisms that may explain the occurrence of this phenomenon. Leptin receptors have
been found in the pituitary [Cai and Hyde 1998] and leptin may directly regulate GH
secretion. Leptin receptors have also been found in the hypothalamus [Mercer et al.
1996] and may regulate the neuropeptides that directly affect GH secretion, GHRH and
SRIH. It has also been shown that leptin inhibits neuropeptide Y (NPY) secretion in the
ARC [Stephens et al. 1995; Schwartz et al. 1996c]. NPY stimulates SRIH and inhibits
GHRH [McCann et al. 1989; Rettori et al. 1990] and therefore inhibits GH, so inhibition
of NPY by leptin would prevent the inhibition of GHRH, thereby causing a stimulation of
GH. The current study does not demonstrate which, if any or all, of these mechanisms
are in effect, but it does show that elevated leptin levels in vivo enhance circulating GH
levels, in support of the first hypothesis.
CHAPTER 4
LEPTIN TREATMENT INCREASES GROWTH HORMONE SECRETION IN
CULTURED GH1 CELLS
Introduction
Growth hormone (GH) is secreted in an episodic pattern from the somatotropic
cells of the anterior pituitary gland. Secretion is under direct hypothalamic control of two
neuropeptides: growth hormone releasing hormone (GHRH) and somatotrope release
inhibiting hormone (SRIH) [Martin and Millard 1986; Millard 1989]. GHRH stimulates
the high amplitude GH secretory episodes. SRIH, on the other hand, is responsible for
prolonged intervals during which little GH secretion occurs. SRIH also regulates GHRH
secretion [Tannenbaum 1994] and the release of both GHRH and SRIH are, in turn,
regulated by many other neuropeptides and neurotransmitters.
One of the many roles of GH is regulation of body composition. GH triggers
lipolysis [Ho et al. 1996] directly at the level of the adipocyte [Fagin et al. 1980; Vikman
et al. 1991]. In obese animals, GH burst amplitudes are reduced [Veldhuis et al. 1991]
but administration of recombinant human GH to rats results in a decrease in body weight.
The exact mechanism of the attenuation of GH in obesity has yet to be elucidated.
Leptin is a hormone produced mainly in adipocytes and which is secreted into the
bloodstream in proportion to the amount of fat present. Leptin crosses the blood-brain
barrier and informs the hypothalamus of the body's energy stores. The hypothalamus
then responds by regulating orexigenic behavior and metabolic rate to maintain body
weight homeostasis.
77
78
Both leptin and GH are related to body composition, and it has been shown that
circulating leptin and GH are intricately intertwined. A previous study in GH-deficient
adults demonstrated a normalization of elevated leptin levels upon GH replacement
[Florkowski et al. 1996, Fisker et al. 1997]. This decrease in leptin most likely occurred
as a consequence of a decrease in body fat due to GH treatment. Another study showed
that leptin treatment induced spontaneous and GHRH-induced GH secretion
[Tannenbaum et al. 1998]. Carro et al. [1997] demonstrated that leptin antiserum
decreased GH amplitude and nadir in rats suggesting that normal levels of leptin are
required for normal GH secretion.
The effects of leptin on GH secretion may occur at the hypothalamic level.
Leptin receptors colocalize with GHRH neurons [Hakansson et al. 1998] in the arcuate
nucleus and are found in the periventricular nucleus [Mercer et al. 1996] where SRIH is
made [Kiyama and Emson 1990]. The regulation of either or both of these two
hypothalamic GH regulatory factors by leptin may result in indirect control of GH by
leptin. In addition, a direct regulation of GH by leptin may occur at the level of the
pituitary. It has recently been shown that leptin and leptin receptors are produced in the
anterior pituitary [Jin et al. 1999; Jin et al. 2000].
In Chapter 3 we demonstrated that leptin induces GH secretion, but there was no
indication of the level of control (hypothalamic or pituitary). The hypothesis of Chapter
4 is that leptin increases GH secretion directly in anterior pituitary cells. To test this
hypothesis, a rat pituitary cell line was used.
79
Methods
GH1 Cells
GH1 cells (American Type Culture Collection, Rockville, MD) are rat pituitary tumor
cells that hypersecrete GH. Cells were grown to 70% confluency in F-12K medium
(ATCC, Rockville, MD) supplemented with 1% non-essential amino acids, 1% Lglutamine, 1% nystatin (Gibco BRL, Life Technologies, Grand Island, NY), and either
10% horse serum and 2.5% fetal bovine serum (FBS) (Gibco BRL, Life Technologies,
Grand Island, NY) or 12.5% charcoal-stripped FBS (Hyclone, Logan, UT). The use of
different serum supplementation is described later in the chapter. Cells were incubated at
37ºC with 5% CO2. GH1 cells were used in passes 42-44.
Plating Cells and Leptin Concentration
In general, cells were plated at 200,000 cells/well in 24-well plates and were allowed
time to adhere, usually 2-3 days, before experimentation. On the day prior to each
experiment, medium was aspirated from each well and discarded. Control or leptinsupplemented (murine leptin, Amgen, Inc., Thousand Oaks, CA) medium was added to
appropriate wells. Leptin was used at a concentration of 100 nM. This concentration of
leptin was chosen because 100 nM seems to be in the upper end of the range of
physiological leptin levels. Experiments were completed as described below.
Five-Day Time -Course
Control or leptin-supplemented medium was added to appropriate wells on each plate.
Supernatant was collected each day for 5 days and frozen at -35ºC until GH RIA. Cells
were collected and analyzed for DNA content for normalization of GH values.
80
Media Experiment
Cells were plated using media and supplements as described above, but with differences
in serum content. Serum-free media or media supplemented with either 10% horse serum
and 2.5% FBS or with 12.5% charcoal-stripped FBS were utilized, as will be described
later in the chapter. Supernatant was collected at 8 and 24 hours and frozen at -35ºC until
GH RIA. Cells were collected and analyzed for DNA content for normalization of GH
values.
RT-PCR for Leptin Receptor
RNA from GH1 cells was collected for RT-PCR amplification of leptin receptor mRNA
as described in Chapter 2. RT-PCR for leptin receptor mRNA was optimized using the
sense and antisense primers designed by Zamorano et al. [1997]. The sense and antisense
primers were 5’-ATGACGCAGTGTACTGCTG and 5’GTGGCGAGTCAAGTGAACCT, respectively and amplified a 357-base pair fragment
of the long form of the leptin receptor.
Western Immunoblot for Leptin Receptor
Leptin receptor protein expression was observed by Western blotting. Briefly, protein
was extracted from hypothalamus, run on a gel, and transferred to a nitrocellulose
membrane. The membrane was then incubated with primary antibody to the long form of
the leptin receptor at a dilution of 1:500 overnight at 4°C and with secondary antibody for
1 hour at room temperature. Detection reagents were added to the membrane, and the
membrane was exposed to radiographic film. The film was developed in a Konica
Medical Film Processor (QX-70).
81
GH Radioimmunoassay
Cell culture media was measured in duplicate for GH using materials supplied by Dr. A.
F. Parlow and the National Hormone and Pituitary Program (NIADDK, Baltimore, MD).
Values were expressed in ng/mL in terms of the NIADDK reference preparation rat GHRP-2. The GH RIA has an assay sensitivity of 1 ng/mL and a range of detection of 1
ng/mL to 320 ng/mL. Leptin at concentrations between 1.6 nM and 160 µM do not
crossreact with the GH assay.
Statistics
Student’s t-test was used to compare GH values between time points and between
treatment groups.
Results
Ob-Rb mRNA and Protein Expression
The band seen at 357 bp represents mRNA for the long-form of the leptin receptor
(Figure 4-1); lanes 3 and 4 are GH1 cells. An additional band, slightly larger, is also
seen. This band is most likely a primer-dimer attached to the fragment of interest, which
can materialize when the MgCl 2 concentration is too high. This extraneous band was
eliminated when the MgCl2 concentration was optimized. For the purposes of this study,
we wanted to know whether or not the leptin receptor was present. Therefore the exact
concentration of leptin receptor mRNA is not required and no normalization with the
housekeeping gene cyclophilin was performed. A Western was completed to determine
if the presence of leptin receptor mRNA correlated to leptin receptor protein. The band
size (in kDa) representing the long-form of the leptin receptor was not revealed in the
82
Western analysis, however, multiple other bands of smaller sizes, also specific to leptin
receptor, were present (data not shown).
Five-Day Time -Course
Over the 5-day time-course, GH1 cells continued to secrete GH (Figure 4-2) and the GH
appeared to be stable in the culture media. Leptin treatment had no effect on GH
secretion.
Media Experiment
In all three serum-supplemented media tested, GH1 cells secreted significantly more GH
by 24 hours than at 8 hours. In media with 10% horse serum and 2.5% FBS (Figure 4-3)
and in serum-free media (Figure 4-4), leptin had no effect at either time point. In media
supplemented with 12.5% charcoal-stripped FBS, leptin significantly increased GH
secretion at 8 hours and tended to increase GH at 24 hour (Figure 4-5).
357 bp
Figure 4-1: Ob-Rb mRNA on GH1 Cells
Lane 1 is the 100 base-pair DNA ladder. Lane 2 is Ob-Rb mRNA in hypothalamus. Lanes 3 and 4 are ObRb mRNA in GH1 cells.
83
Figure 4-2: Five-Day Time-Course
Over the 5-day time-course, GH1 cells continually secreted GH (cumulative results over time). Leptin
treatment had no effect on GH secretion (t-test, n=6 each). Cells were plated in media supplemented with
10% horse serum and 2.5% FBS.
Figure 4-3: Media Supplemented with 10% Horse Serum and 2.5% FBS
Significantly more GH was secreted at 24 hours (n=6) than at 8 hours (t-test, p<0.05, n=6). Leptin had no
effect at either 8 (n=6) or 24 hours (n=6).
84
Figure 4-4: Serum-Free Media
Significantly more GH was secreted at 24 hours (n=6) than at 8 hours (t-test, p<0.05, n=6). Leptin had no
effect at either 8 (n=6) or 24 hours (n=6).
Figure 4-5: Media Supplemented with 12.5% Charcoal Stripped FBS
Significantly more GH was secreted at 24 hours (n=6) than at 8 hours (t-test, * = p<0.05, n=6). Leptin
treatment significantly increased GH secretion at 8 hours (t-test, ** = p<0.05) and the effects were nearly
significant (p=0.07) at 24 hours (n=6 each).
85
Discussion
Leptin has been shown to influence GH secretion indirectly through regulation of
hypothalamic neuropeptides [Mercer et al. 1996; Hakansson et al. 1998]. In addition, a
direct regulation of GH by leptin may occur at the pituitary level; it has recently been
shown that leptin and leptin receptors are produced in the anterior pituitary of humans
[Jin et al. 1999] and rodents [Jin et al. 2000]. The current study shows that long-form
leptin receptor mRNA is present in GH1 cells. When analyzed for protein expression, the
long form of the leptin receptor was not seen, but many other isoforms were. Until
recently, it was thought that only the long form of the leptin receptor was biologically
active; however, it is now known that the short forms can also transduce signals
[Bjorbaek et al. 1997; Murakami et al. 1997; Yamashita et al. 1998].
GH1 cells secrete continuously over time in the absence of a stimulus. In the
time-course presented here, GH levels increased consistently over 5 days indicating that
GH was stable in the media over time. Leptin treatment had no effect on GH secretion on
any of the 5 days. However, these cells were plated in media supplemented with 10%
horse serum and 2.5% FBS. After completing these studies, it was discovered that using
serum-supplemented media could interfere with the actions of leptin on these cells.
To further investigate the effects of serum on leptin treatment, a study was
undertaken in which media with different serum contents were utilized. As was seen in
the time-course, cells in medium supplemented with 10% horse serum and 2.5% FBS
continued to secrete GH over time, but leptin had no effect. There are several potential
explanations for this. There may be leptin-binding proteins in the serum that prevent
leptin from binding to its cells surface receptors. Conversely, there may be some factor
86
in the serum that antagonizes leptin actions. Additionally, the serum may contain
elements that degrade or alter leptin, making it biologically inactive.
To eliminate these potentially confounding effects of serum on leptin, serum-free
medium was utilized. Again, cells secreted significantly more GH by 24 hours than at 8
hours, but leptin had no effect at either time point. Leptin did, however, tend to decrease
GH at 24 hours. This seems contrary to the hypothesis that leptin stimulates GH
secretion, however, leptin and GH are substances that may stick to the plastic 24-well
plates. In media supplemented with serum, the serum will coat the plates and prevent
leptin and GH from sticking. The results of the current study indicated that, in serumfree media, leptin tended to decrease GH secretion. However, it is probable that leptin
was bound to the plate and therefore could not produce an effect. In addition, GH may
have also adhered to the plate, possibly explaining why there appears to have been less
GH secreted in Figure 4-4 than in Figure 4-3.
In an attempt to eliminate the uncertainty of the effects of either serumsupplemented or serum-free media on the actions of leptin on GH1 cells, charcoalstripped FBS was utilized. Charcoal stripping reduces the levels of many hormones,
growth factors, and steroids. In media supplemented with charcoal-stripped FBS, leptin
treatment significantly increased GH secretion at 8 hours, in support of the original
hypothesis. Leptin also tended to increase GH secretion at 24 hours, indicating that leptin
lost some of its effectiveness over time. With continuous leptin treatment, leptin
receptors on GH1 cells may be downregulated. In the future, an 8-hour time-course and a
leptin dose-response should be completed. In addition, leptin receptor mRNA
concentration could be measured in the normal and leptin-resistant states.
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The GH data reported was normalized to DNA content. It was found that leptin
treatment of GH1 cells in charcoal-stripped media decreased DNA content at both 8 and
24 hours, indicating that cell proliferation was inhibited. These results agree with those
in which leptin treatment decreased proliferation in GH3 cells [Jin et al. 2000]. The GH3
cell line was initiated from the same primary culture from which the GH1 cells were
initiated, two passages later. When we analyzed raw GH data, leptin had no effect (data
not shown) but when we analyzed GH data normalized to DNA, leptin stimulated GH
secretion. Taken together, these results suggest that leptin inhibits cell proliferation but
stimulates GH secretion from the cells that are present. GH1 cells are tumor cells, so any
extrapolation to in vivo physiology is uncertain; however, perhaps these results indicate
that leptin stimulates GH while decreasing cell proliferation as a self-limiting process.
This should be studied in the future in primary pituitary cells.
In vivo, serum is not charcoal-stripped, nor is there a serum-free option, so it may
be argued that the methods utilized in this chapter are not indicative of what happens in
whole animal physiology. In animals, serum is present that contains leptin-binding
proteins. However, it is possible that these proteins have a lower affinity for leptin than
leptin receptors, so when bound leptin reaches the desired destination, the binding
proteins would release leptin making it available to act at its receptor.
In summary, the studies presented here indicate that GH1 cells express leptin
receptor mRNA and that leptin stimulates GH secretion from these cells. These results
support the first hypothesis of this dissertation.
CHAPTER 5
LEPTIN RESISTANCE IS ASSOCIATED WITH HYPOTHALAMIC RECEPTOR
mRNA AND PROTEIN DOWNREGULATION
Introduction
The ob gene expresses the "obese protein," leptin, which is secreted from
adipocytes and circulates in the plasma. Leptin has been implicated as one of the signals
that is sensed by the hypothalamus to regulate feeding and energy expenditure thereby
maintaining body weight homeostasis [Zhang et al. 1994; Campfield et al. 1995; Halaas
et al. 1995; Pelleymounter et al. 1995; Halaas et al. 1997]. Leptin is expressed primarily
in adipocytes, including those of epididymal, parametrial, abdominal, perirenal, and
inguinal fat pad origin [Maffei et al. 1995a; McGregor et al. 1996] and, to a much lesser
extent in brown adipose tissue [Maffei et al. 1995a]. Moreover, leptin is secreted in
proportion to adipose mass [Considine et al. 1996b; Maffei et al. 1995b].
Leptin receptors are located in many peripheral tissues [Cioffi et al. 1996; Lee et
al. 1996; Tanizawa et al. 1997] and the central nervous system (CNS). In the CNS, leptin
receptors are especially abundant in areas of the hypothalamus that are implicated in
body weight regulation: arcuate, ventromedial, paraventricular, and ventral
premammillary nuclei [Mercer et al. 1996].
Many splice variants of the receptor have been found. Most isoforms have short
intracellular domains, some of which are thought to regulate transport of leptin into the
brain. Other roles for these isoforms have yet to be elucidated. Another type of splice
variant has a longer intracellular domain. This is the variant through which leptin exerts
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a biological effect. Recently, a soluble isoform of the leptin receptor has been
characterized which has been proposed to regulate leptin bioactivity [Liu et al. 1997]. A
mutation of any of these isoforms could result in resistance to leptin, as is seen in the
obese fa/fa rat and db/db mouse.
Mice with mutations of the ob gene have reduced levels of circulating leptin and
exhibit hyperphagia and obesity. When these animals are given exogenous leptin, their body
weight is significantly reduced. Unfortunately, these findings are not easily extrapolated to
human obesity. Many obese humans exhibit dramatically elevated levels of circulating leptin
[Zhang et al. 1994; Maffei et al. 1995b] that can result in an inability of the hypothalamus to
detect leptin. When this occurs, the result is increased feeding, added fat mass, and further
elevated levels of circulating leptin. Failure of the hypothalamus to detect leptin, or leptin
resistance, may occur for one of several reasons. First, leptin may fail to cross the bloodbrain barrier. Secondly, the hypothalamic receptors may be mutated or downregulated.
Alternatively, downstream signaling may be inhibited. Although investigators are not all of
the same opinion, most investigators believe in the development of a leptin-resistant state
with the progression toward obesity in both humans [Caro et al. 1996a] and animals
[Frederich et al. 1995a; Maffei et al. 1995b; Van Heek et al. 1997]. The current study
demonstrates that resistance to the anorectic and GH-regulating effects of leptin develops in
conjunction with downregulation of hypothalamic receptor in male Long-Evans rats.
Methods
Animals
Thirty-six male Long-Evans rats (150-174 g, 42-45 days, Harlan Sprague-Dawley) were
obtained from Harlan Sprague-Dawley and housed individually under standard lighting
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and temperature conditions. The rats were fed normal chow, which, along with water,
was available ad libitum. The animals were randomly divided into 3 leptin-treated
groups. Body weight and food intake were measured daily for the duration of the study.
Groups
Three groups of animals (12 per group) were implanted subcutaneously with Alzet
osmotic pumps (Model 2ML4, Alza Scientific Products, Palo Alto, CA) that deliver their
contents at a rate of 2.5 µL/hr for 28 days. The contents of the pumps consisted of sterile
PBS (control), leptin at a dose of 0.1 mg/kg/day (low dose), or leptin at a dose of 0.5
mg/kg/day (high dose). The lower dose was chosen to mimic physiological leptin levels
in obese rats (10-25 ng/mL) while the higher dose was chosen to test for resistance in the
event that it was not seen at the lower dose. The murine leptin used in this study was
provided by Amgen, Inc. (Thousand Oaks, CA).
Serum Measurements
Blood samples were collected by cardiac puncture on the day of implantation (day 1) and
on day 15; trunk blood was collected at sacrifice (day 29) for determination of leptin,
insulin-like growth factor-1 (IGF-1), insulin, glucose, and triglycerides. All samples
were collected in the morning between 8 AM and 12 PM. Leptin was measured with a
rat leptin radioimmunoassay (RIA) kit (Linco Research, Inc., St. Charles, MO). This kit
measures both rat and mouse leptin with an assay sensitivity of 0.5 ng/mL and a range of
detection from 0.5 ng/mL to 50 ng/mL. IGF-1 was extracted from blood by the
acid/ethanol procedure and measured by RIA as previously described [Grant et al. 1986].
The IGF-1 RIA has an assay sensitivity of 0.1 ng/mL and a range of detection from 0.1
ng/mL to 20 ng/mL. Insulin was measured with a rat insulin RIA kit (Linco Research,
Inc., St. Charles, MO) with an assay sensitivity of 0.1 ng/mL and a range of detection
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from 0.1 ng/mL to 10 ng/mL. Glucose was measured using a YSI 1500 Sidekick Glucose
Analyzer (Yellow Springs Instrument Company, Inc., Yellow Springs, OH).
Triglycerides were measured using a kit from Sigma (Sigma Diagnostics, St. Louis, MO).
Leptin Challenge Test Pilot Study
Dose and route of leptin administration were determined in a pilot study prior to the
actual resistance study. Naïve rats were food deprived for 24 hours then given an
intraperitoneal (IP) or subcutaneous (sc) leptin or PBS injection, utilizing different doses
of leptin. In two studies, leptin was given IP at either 1.5 or 2.8 mg/kg. In another study,
30 mg/kg leptin was given sc. Food was returned after each injection, and food intake
was measured at 24 hours.
Leptin Challenge Test
A test of leptin resistance was performed on day 21. The animals were food deprived for
24 hours prior to the test. The leptin challenge test consisted of a sc bolus of murine
leptin (30 mg/kg) which was given to half of the animals in each treatment group (leptin
challenge). The other half of the animals in each group received a bolus of PBS for
control. Food (Purina rat chow #5001) was returned to the animals one hour postinjection, allowing the animals time to recover following the injection. Food intake was
then measured at 4 hours and again at 24 hours.
RT-PCR for Leptin Receptor
RT-PCR for leptin receptor mRNA was optimized using the sense and antisense primers
from Zamorano et al. [1997]. The sense and antisense primers were 5’ATGACGCAGTGTACTGCTG and 5’-GTGGCGAGTCAAGTGAACCT, respectively
and amplified a 357-base pair fragment of the long form of the leptin receptor. Optimal
conditions were identified as 1 mM MgCl 2, 64°C annealing temperature, and 30 cycles
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following an initial denaturation phase and followed by a final elongation phase. The
identity of RT-PCR product was confirmed using the internal probe 5’TGCAGCTGAGGTATCACAGG in Southern blot analysis (ECL, Amersham Pharmacia
Biotech, Piscataway, NJ). The amount of leptin receptor mRNA was normalized with the
housekeeping gene cyclophilin. The cyclophilin sense and antisense primers used were
5'-GGGAAGGTGAAAGAAGGCAT and 5'-GAGAGCAGAGATTACAGGGT,
respectively and amplified a 210-base pair fragment [Zamorano et al. 1997]. The same
conditions were used for cyclophilin as were used for the leptin receptor.
Western Immunoblot for Leptin Receptor
Leptin receptor protein expression was measured by Western blotting. Briefly, protein
was extracted from hypothalamus, run on a gel, and transferred to a nitrocellulose
membrane. The membrane was then incubated with primary antibody at a dilution of
1:500 overnight at 4°C and with secondary antibody for 1 hour at room temperature.
Detection reagents were added to the membrane, and the membrane was exposed to
radiographic film. The film was developed and analyzed with Molecular Analysis
Software (Hercules, CA). The results were normalized according to the amount of
protein loaded, and the results are expressed as a ratio of sample:intra-gel control.
Statistics
Body weight, food intake, IGF-1 levels, and leptin values were analyzed by two-way
repeated measures ANOVA followed by Student-Newman-Keuls Multiple Comparisons.
Comparisons between the leptin challenge and control groups were by Student's t-test for
each pilot study. Two-way ANOVA and Duncan's Multiple Comparisons were used to
analyze food intake in response to the leptin challenge. One-way ANOVA followed by
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Tukey’s or Dunn's Multiple Comparisons was used to analyze insulin, glucose, and
triglyceride, and leptin receptor mRNA and protein expression data.
Results
Leptin
Leptin levels were significantly elevated in the low-dose implant group after 4 weeks of
treatment as well as in the high-dose implant group after 2 and 4 weeks of treatment
compared to control (Figure 5-1). At time of implantation, there were no differences in
serum leptin levels among the groups. Within the high-dose group, leptin levels were
significantly elevated at weeks 2 and 4 compared to week 0.
Body Weight
Rats in the control group weighed significantly more than rats in the high-dose group at
day 7 and from day 9 through the remainder of the study (Figure 5-2). Body weight
between the two leptin-treated groups was significantly different at days 12, 15-19, and
21-29. The differences in body weight between the low-dose group and the controls
never reached statistical significance although there was a trend for the low-dose rats to
weigh less than the control rats. All 3 groups experienced a drop in body weight
following 24-hour periods of food deprivation, indicated by discontinuations of graph
lines.
Food Intake
Rats in the high-dose group consumed significantly less rodent chow than controls on
days 4-9, 16, and 23 (Figure 5-3). Rats in the low-dose group also ate significantly less
than controls on days 4-6 and 23. The difference in food intake in the two leptin-treated
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groups was significantly different at days 4-9. Periods of food deprivation are indicated
by discontinuations of graph lines.
Leptin Challenge Test Pilot Studies
Neither the 1.5 mg/kg nor the 2.8 mg/kg dose of leptin administered IP was effective at
reducing food intake at 24 hours. Leptin given sc at a dose of 30 mg/kg reduced 24-hour
food intake significantly compared to control (Figure 5-4). This dose and route of leptin
administration (leptin challenge) was used in the leptin challenge test.
Leptin Challenge Test
At 24 hours, the rats in the control implant group ate significantly less when challenged
with leptin compared to their respective controls whereas the leptin challenge had no
effect in rats treated with either dose of leptin (Figure 5-5). In addition, there were no
differences in food intake among the treatment groups due to the leptin challenge, but
food intake of the low- and high-dose leptin groups were significantly lower than controls
following PBS control injection.
Leptin Receptor mRNA in Hypothalamus
In both leptin-treated groups, leptin receptor mRNA was significantly reduced when
compared to control (Figure 5-6). Leptin receptor mRNA was normalized to the
housekeeping gene cyclophilin.
Leptin Receptor Protein Expression in Hypothalamus
The high-dose leptin-treated group had significantly reduced leptin receptor protein
expression compared to control (Figure 5-7). The low-dose leptin group also tended to
have less protein expression compared to control, but the results were not significant.
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IGF-1 Values
At week 2, the high-dose leptin group had significantly reduced IGF-1 values compared
to week 0 (Figure 5-8). At week 4, both the control and low-dose leptin groups had
significantly elevated IGF-1 values compared to week 0. Also at week 4, IGF-1 was
significantly lower in the high-dose leptin group than in either of the other two groups.
Hormonal and Metabolic Measures
Measures of hormonal and metabolic state including insulin, glucose, and triglycerides
are reported in Table 5-1. At both weeks 2 and 4, triglycerides were significantly
attenuated in the high-dose leptin group compared to the other two groups. At week 4,
glucose was significantly reduced in the high-dose group compared to the other two
groups. Also at week 4, insulin was significantly lower in both the low- and high-dose
groups compared to control.
Figure 5-1: Leptin Levels Indicate Proper Pump Activity
Serum leptin levels were significantly elevated (* = p<0.05 vs. control) in the low dose treatment group
(n=12) compared to control (n=12) and in the high-dose treatment group (n=12) at weeks 2 and 4 compared
to control (2-way repeated measures ANOVA). Serum leptin levels were significantly higher (# = p<0.05
vs. week 0) in the high-dose treatment group at weeks 2 and 4 compared to week 0.
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Figure 5-2: Body Weight is Dose-Dependently Decreased by Leptin Treatment
There were no significant differences in body weight between the control (n=12) and low dose treatment
groups (n=12) although the low dose group tended to weigh less for the duration of the study (2-way
repeated measures ANOVA). Body weight in the high dose treatment group (n=12) was significantly
decreased (p<0.05) at days 12, 15-19, and 21-29 compared to the low dose group and at days 7 and 9-29
compared to control (2-way repeated measures ANOVA). Discontinuations of graph lines follow 24-hour
periods of food deprivation resulting in erroneous decline in body weight.
Figure 5-3: Food Intake is Initially Decreased by Leptin; Ultimately Resistance Develops
Food intake was significantly inhibited (p<0.05) in the low dose treatment group (n=12) compared to
control (n=12) at days 4-6 and 23 (2-way repeated measures ANOVA) and in the high dose treatment group
(n=12) compared to control at days 4-9, 16, and 23. Food intake was significantly different (p<0.05)
between the two treatment groups at days 4-9 (2-way repeated measures ANOVA). Discontinuations of
graph lines represent 24-hour periods of food deprivation.
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Figure 5-4: Leptin Challenge Test Pilot Study
In the pilot study which used a leptin challenge of 30 mg/kg administered subcutaneously (sc), leptin
significantly reduced food intake (p<0.05, Student's t-test) compared to control. Neither the 1.5 mg/kg nor
the 2.8 mg/kg intraperitoneal (IP) leptin challenge was effective at reducing food intake. Each group, n=6.
Figure 5-5: Leptin Challenge Test
At 24 hours, food intake was significantly inhibited (p<0.05) by leptin challenge (n=6) compared to
respective controls (n=6) within the treatment control group (2-way ANOVA). There were no differences
in food intake as the result of leptin challenge (n=6) in either leptin-treated group compared to respective
controls (n=6). Food intake in the control treatment group was significantly higher than in their low- and
high-dose leptin counterparts following the PBS control injection.
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Figure 5-6: Leptin Receptor mRNA in Hypothalamus
In both the low-dose (n=12) and high-dose (n=10) leptin groups, leptin receptor mRNA was significantly
reduced in hypothalamus (p<0.05, 1-way ANOVA) compared to control (n=12). Leptin receptor mRNA
was normalized to the housekeeping gene cyclophilin.
Figure 5-7: Leptin Receptor Protein Expression in Hypothalamus
The high-dose leptin-treated group (n=9) had significantly reduced leptin receptor protein expression
compared to control (p<0.05, 1-way ANOVA, n=12). The low-dose leptin group (n=9) also tended to have
less protein expression compared to control, but the results were not significant.
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Figure 5-8: IGF-1 Values
At week 2, the high-dose leptin group had significantly reduced IGF-1 values compared to week 0 (p<0.05,
2-way repeated measures ANOVA). At week 4, both the control and low-dose leptin groups had
significantly elevated IGF-1 values compared to week 0. Also at week 4, IGF-1 was significantly lower in
the high-dose leptin group than in either of the other two groups. N=8-12.
Table 5-1: Hormonal and Metabolic Measures
Week 0
Insulin
Glucose (mg/dL)
(ng/mL)
Control
N=10-12
1.96 ± 0.24
177 ± 4.93
Low Leptin
N=8-12
1.73 ± 0.24
176 ± 5.05
High Leptin
N=7-12
2.28 ± 0.24
164 ± 4.16
Week 2
Control
Low Leptin
High Leptin
Glucose (mg/dL)
N=9-10
N=3-8
N=3-8
Insulin
(ng/mL)
1.04 ± 0.18
0.41 ± 0.11
0.81 ± 0.13
Glucose (mg/dL)
N=12
N=12
N=8-12
Insulin
(ng/mL)
1.72 ± 0.16
1.17 ± 0.12 *
0.77 ± 0.15 *
Week 4
Control
Low Leptin
High Leptin
168 ± 3.36
162 ± 3.34
162 ± 1.53
158 ± 3.29
156 ± 2.42
144 ± 2.75 **
Triglycerides
(mg/dL)
103.20 ± 8.10
110.09 ± 9.08
107.31 ± 8.25
Triglycerides
(mg/dL)
81.37 ± 5.88
72.99 ± 8.22
47.66 ± 4.59 **
Triglycerides
(mg/dL)
122.16 ± 7.00
98.77 ± 9.89
57.73 ± 9.04 **
Values are given as mean ± SEM. * = p<0.05 vs. control. ** = p<0.05 vs. control and low leptin. 1-way
ANOVA.
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Discussion
At time of osmotic pump implantation (week 0), there were no differences in
serum leptin levels among the different treatment groups. At week 2, the high-dose
group had significantly elevated leptin levels compared to control and to the low-dose
group. At week 4, both the high-dose and the low-dose groups had significantly elevated
leptin levels compared to control. In addition, serum leptin levels were significantly
elevated in the high-dose leptin-treated group at weeks 2 and 4 compared to week 0.
Taken together, these results indicate that the osmotic pumps were effective at delivering
their contents and the murine leptin was stable. Additionally, leptin has been shown to be
structurally stable after 4 weeks at 37°C (MaryAnn Pelleymounter, Neurocrine
Biosciences, Inc., personal communication). It was previously shown that, within 24
hours of leptin withdrawal, body weight normalizes immediately (MaryAnn
Pelleymounter, Neurocrine Biosciences, Inc., personal communication). A normalization
of body weight was not observed in the current study indicating that leptin continued to
be effective over time; in addition to its stability, leptin maintained its biological activity.
With continuous infusion of drug it would be expected that, once established,
steady state serum levels would be maintained over time. On the contrary, leptin levels
in these rats continued to increase throughout the study period, indicating that steady state
was never achieved. One potential mechanism by which this occurred could have been a
saturation of clearance or a downregulation of "clearance" receptors (Ob-Ra on kidney);
however, it was previously shown that, even when pharmacological doses of leptin are
administered, elimination pathways are not saturated [Cumin et al. 1996; Hill et al.
1998]. It is unknown if the doses used in this dissertation exceeded the pharmacological
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doses used in saturation or elimination studies. The more likely explanation of lack of
steady state leptin levels is the result a pseudo-two-compartment model of drug infusion.
Upon implantation, cysts rapidly form around the osmotic pumps. As expected, some of
the leptin being released from the pump gets into the bloodstream. In addition, some of
the leptin may also be sequestered by the cyst. Over time, as the cyst gets saturated with
leptin, leptin would be released from the cyst as well as from the pump. This would
result in higher serum leptin levels over time, as is seen in the current study.
It is generally believed that with leptin treatment food intake is attenuated. This
was seen in the first two weeks of our study in a dose-dependent manner: the animals
treated with high-dose leptin ate significantly less than those treated with low-dose leptin,
which in turn ate less than controls. During the final two weeks of the study, there were
virtually no differences in food intake among the groups indicating that the rats in each
treatment group became resistant to the long-term appetite regulating effects of leptin.
This loss of effect of leptin treatment on food intake over time was shown previously by
us (unpublished observations) and others [Pelleymounter et al. 1995]. The mechanism by
which the anorectic effects of leptin were lost is likely the downregulation of
hypothalamic leptin receptor mRNA and protein that occurred as a pharmacological
response to the higher levels of circulating leptin.
It appeared that the leptin-treated rats began to develop resistance to the anorectic
effects of leptin approximately half way through the study, so we performed a leptin
challenge test at week 3. The leptin challenge test was a one-time sc bolus of a dose of
leptin high enough to attenuate 24-hour food intake in control rats. Within each
treatment group, half of the animals received the leptin challenge and the other half
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served as controls and received an injection of PBS. As expected within the control
treatment group, the leptin challenge acted to decrease food intake at 24 hours.
Alternatively, the rats within the leptin treatment groups did not respond to the leptin
challenge; there were no significant differences in food intake in either treatment group at
either time point following the leptin challenge compared to their respective controls.
These results, together with the food intake results, indicate that continuous exogenous
leptin treatment results in the development of resistance to the appetite regulating effects
of leptin by the 2nd or 3rd week of treatment. Additionally, in the 4-hour leptin
challenge test in the low-dose leptin group, there was a statistically nonsignificant
increase in food intake in response to the leptin challenge. By 24-hours, however, there
was no detectable effect of leptin on food intake in the chronically treated groups.
An important observation with this test must be discussed. The control rats in the
chronic PBS treatment group ate significantly more than the control rats in either chronic
leptin treatment group. These results suggest that, although resistance to the appetiteregulating effects of leptin was developing in the animals treated chronically, the longterm leptin treatment resulted in some sort of ceiling effect is seen in which the rats
simply cannot eat as much as untreated rats. In addition, the leptin-treated animals were
eating the same amount as the untreated animals at this time point.
Leptin treatment decreases or attenuates increases in body weight. At the
beginning of the current study, leptin treatment dose-dependently decreased body weight,
which corresponded with the inhibitory effects on food intake. After the second week,
leptin did not cause further decrease in body weight and the rate of growth was similar in
all three groups indicating that “catch-up” growth was prevented. Unlike the food intake
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effects of leptin, the animals did not become resistant to the body weight regulating
effects of leptin. Our body weight and food intake results correspond to those from a
study in which mice, after receiving daily leptin injections, experienced a normalization
of the initially reduced food intake [Pelleymounter et al. 1995]. In that study, the
reduction in body weight was never normalized. In this chapter, the body weight never
normalized.
IGF-1 has a relatively long half-life and is secreted more continuously than GH
and therefore is a useful indicator of GH secretory status [Blum et al. 1990]. In the
control and low-dose leptin groups, IGF-1 levels were significantly elevated by week 4.
This may be explained in the controls due to normal growth. The results of the IGF-1
seen in the low-dose leptin group support the first hypothesis of the dissertation. It may
be expected, however, that since the low-dose group had much higher leptin levels than
the control group, the low-dose group should also have significantly elevated IGF-1
values compared to the control group. This was not seen, perhaps indicating that
resistance to the GH-stimulating effects of leptin were beginning to be lost. In support of
this suggestion, hypothalamic leptin receptor protein expression was apparently
downregulated, although the effects were not significant. In the high-dose leptin group,
IGF-1 values declined over time and were lower than IGF-1 in either of the other two
groups at week 4. The results of Figure 5-8 indicate that, at this length of time of pump
implantation, the lower dose of leptin was not sufficient and higher dose of leptin was
sufficient to cause resistance to the GH-stimulating effects of leptin.
There are different mechanisms by which food intake and body weight are
regulated by leptin, including central alterations in metabolic rate and energy expenditure
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as well as direct effects on fat cells. Neither metabolic rate/energy expenditure nor fat
content were measured in this study. We did, however, measure other indices of
metabolic action and found decreases in serum insulin, glucose, and triglyceride levels in
the high-dose leptin group as well as a decrease in insulin in the low-dose group by week
4. It should be pointed out the basal glucose in the control rats is elevated, probably due
to the stressful sampling technique. The results of Table 5-1 indicate that leptin retained
its ability to affect metabolic parameters even though its ability to alter food intake was
lost. The results of the food intake, leptin challenge, IGF-1, and metabolic studies
indicate that the behavioral and metabolic aspects of leptin resistance occur by different
mechanisms and at different levels of circulating leptin.
One type of leptin resistance occurs as the result of a mutation of the leptin
receptor. There are various types of leptin receptor mutations that result in obesity in
rodents [Flier and Elmquist 1997] as well as in humans [Clement et al. 1998]. Leptin
resistance can also develop over time as opposed to being the result of a mutation. In a
recent study in obese mice with peripheral resistance to leptin, it was shown that central
leptin administration had the desired effect of reduction of food intake and body weight
[Van Heek et al. 1997]. These results demonstrate that leptin resistance may occur if
leptin progressively loses its ability to cross the blood-brain barrier. Transport of leptin
across the blood-brain barrier is unidirectional from blood to brain and occurs by a
saturable process [Banks et al. 1996; Caro et al. 1996a] via short-form leptin receptors.
Perhaps in response to the slowly developing hyperleptinemia that occurs with increasing
body weight, there is a down-regulation of these leptin transporters [Caro et al. 1996a]. It
must be noted, however, that several of the hypothalamic nuclei where leptin exerts many
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of its biological effects are circumventricular organs, located near fenestrations of the
blood-brain barrier.
Other forms of leptin resistance can occur in a pharmacokinetic manner, for
example if there is a problem with distribution, delivery, metabolism, and/or elimination.
Leptin resistance can also occur in a pharmacodynamic manner, with receptor
downregulation or disruption of signal transduction. More than likely, there are multiple
potential mechanims for the development of leptin resistance. It has been shown there
are varying degrees of leptin resistance among different strains of obese mice [Halaas et
al. 1995] and there are likely to be variations of leptin resistance in obese humans as well.
There are multiple complex CNS pathways through which leptin produces bioactivity
[Elmquist et al. 1998], and a problem with any one of them can result in a perturbation of
body weight homeostasis.
An important part of the mechanism(s) by which leptin resistance occurred in our
study incorporated downregulation of hypothalamic leptin receptor mRNA and protein
expression. It is a well-known pharmacological phenomenon that elevated levels of
agonist, in this case circulating leptin, result in downregulation of receptor to prevent
excessive activity. When this occurrence is chronic, there develops a state of resistance
to the agonist. This response is analogous to insulin resistance seen in hyperinsulinemia.
There is a strong historical precedence for agonist-induced receptor downregulation and
our findings on leptin receptor downregulation support this model of resistance. This is
an important and heuristic finding.
In conclusion, we have demonstrated the development of resistance to the
anorectic and GH-regulating effects of leptin in conjunction with downregulation of
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hypothalamic leptin receptor mRNA and protein expression. These results support the
second hypothesis of this dissertation.
CHAPTER 6
LEPTIN'S EFFECTS ON FOOD INTAKE AND BODY WEIGHT ARE
DIFFERENTIALLY ATTENUATED IN RATS FED A LOW-FAT OR HIGH-FAT
DIET
Introduction
Leptin is a hormone secreted primarily by adipose tissue [Maffei et al. 1995a;
McGregor et al. 1996]. It travels in the bloodstream, actively crosses the blood-brain
barrier, and binds to its receptors in the hypothalamus to inform the brain of the body’s
energy stores. Leptin levels increase in parallel with body fat [Considine et al. 1996b;
Maffei et al. 1995b]. When the brain senses this elevation of leptin levels, the response is
decreased feeding and heightened metabolic rate [Zhang et al. 1994; Campfield et al.
1995; Halaas et al. 1995; Pelleymounter et al. 1995; Halaas et al. 1997]. Conversely, the
absence of leptin that is seen in the ob/ob mouse or defective leptin signaling that is seen
in the db/db mouse and the fa/fa rat results in obesity due to hyperphagia and a low
metabolic rate. Exogenous leptin administered to leptin-deficient obese animals results in
a lowered body weight due to the restoration of normal feeding patterns and metabolism
[Campfield et al. 1995; Halaas et al. 1995; Pelleymounter et al. 1995]. In addition, the
reduction in body weight is observed when leptin is administered to leptin-producing
animals of normal body weight.
Leptin’s behavioral effects are attenuated in animals fed a highly palatable diet
[Campfield et al. 1995; Frederich et al. 1995a; Masuzaki et al. 1995; Van Heek et al.
1997; Widdowson et al. 1997] even though leptin is increased. It would be expected that
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108
the elevated leptin levels would cause a decrease in food intake and body weight
[Campfield et al. 1995], but conversely, food intake and body weight were elevated in
these studies [Frederich et al. 1995a; Masuzaki et al. 1995; Van Heek et al. 1997;
Widdowson et al. 1997]. Studies completed to date examined elevated leptin levels that
originated from endogenous sources and focused on the effects of high-fat diets. The
current study examines the effects of exogenously elevated serum leptin and was
undertaken to test the hypothesis that induced hyperleptinemia, due to exogenous leptin
administration, also fails to be effective in rats fed a high-fat, high-calorie diet.
Furthermore, the study was designed to ascertain if the same would be observed in rats
fed a low-fat, high-calorie diet. The same diets were used in this study as were used in
Chapter 3, but the animals were fed for a longer period of time before experimentation to
allow the development of resistance. In addition, this study utilizes continuous delivery
of leptin via osmotic minipumps, as was used in Chapter 5. In this chapter, leptin was
given for four weeks at a dose between the two doses used previously.
Methods
Animals and Diets
Twenty-two male Long-Evans rats (90-95 days, 350-450 g) were obtained from HarlanSprague Dawley and housed individually under standard temperature and lighting
conditions. The animals were randomly divided into three groups (n=7 or 8), each
receiving varied diets ad libitum: normal rat chow (5% fat, 3.0 kcal/g, Purina Mills, Inc.,
Richmond, VA), high-fat, high-calorie diet (23% fat, 3.7 kcal/g, PJ Noyes Company,
Inc., Lancaster, NH), or low-fat, high-calorie diet (2% fat, 3.7 kcal/g, PJ Noyes
Company, Inc., Lancaster, NH). It should be noted that the low-fat diet had a calorie
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content equivalent to that of the high-fat diet, both of which were higher in calories than
the control diet. Water was available to all groups ad libitum. Body weight and food
intake were measured daily for the duration of the study. Rats were sacrificed by
decapitation and the weights of brains, pituitaries, hearts, testes, kidneys, livers, and
adrenals were measured. Hypothalami were snap frozen in liquid nitrogen and stored at
-90°C until further analysis.
Osmotic Pumps, Blood Sampling, and Body Temperature
After 50 days on the various diets, half the rats in each group were implanted
subcutaneously with leptin-filled Alzet osmotic mini-pumps (Alza Scientific Products,
Palo Alto, CA) for the continuous delivery of leptin (0.25 mg/kg/day; 5 µg/hour) over a
14-day period; the other half were implanted with pumps delivering vehicle (control).
Murine leptin was provided by Amgen, Inc. (Thousand Oaks, CA). Blood samples were
collected from the tail vein prior to pump implantation (week 0) and at week 1 and trunk
blood was collected at week 2 for measurement of leptin and IGF-1. All samples were
collected in the morning between 8 AM and 12 PM. Blood was centrifuged and serum
was stored at -20°C until analysis. Body temperature was measured at weeks 0, 1, and 2
with an anal thermistor (Cole Parmer, Vernon Hills, IL).
Leptin and IGF-1 Radioimmunoassays
Serum leptin was measured with a rat leptin RIA kit (Linco Research, Inc., St. Charles,
MO). This kit measures both rat and mouse leptin with an assay sensitivity of 0.5 ng/mL
and a range of detection from 0.5 ng/mL to 50 ng/mL. IGF-1 was extracted from blood
by the acid/ethanol procedure and measured by RIA as previously described [Grant et al.
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1986]. The IGF-1 RIA has an assay sensitivity of 0.1 ng/mL and a range of detection of
0.1 ng/mL to 20 ng/mL.
Statistics
Body weight, food intake, and leptin values were analyzed by two-way repeated
measures ANOVA followed by Student-Newman-Keuls Multiple Comparisons. Twoway ANOVA was used to analyze temperature each week and organ weights, and IGF-1
values followed by Duncan's Multiple Comparisons.
Results
Effects of Diet on Food Intake
Figure 6-1 shows absolute food intake for the duration of feeding of the three different
diets. Since there are differences in kcal and percent fat in each diet, food intake is
reported in grams of food (panel A), kcal (panel B), and grams of fat (panel C) ingested.
The normal chow rats ingested more grams of food than the other two groups for a
majority of the study, but there were no differences in the intake of kcal among the three
groups. As expected, the high-fat fed rats ingested the highest amount of fat while the
low-fat rats ingested the lowest.
Leptin
Basal serum leptin levels were not different among the PBS-implanted groups (Figure 62). As expected, leptin levels did not change significantly over the 2-week implantation
time in rats of any diet group implanted with control pumps. In the rats implanted with
leptin-filled osmotic pumps, serum leptin levels increased significantly in each of the
groups over the two-week period.
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Effects of Diet and Leptin on Food Intake
As expected, leptin treatment decreased food intake (measured in grams of food) in rats
fed normal chow for most of the study (Figure 6-3 panel a). Unexpectedly, on days 7, 12,
and 13, leptin treated actually increased food intake in rats fed the high-fat diet (Figure 63 panel b). There was no effect of leptin on food intake in rats fed the low-fat diet
(Figure 6-3 panel c). Food intake was also shown as grams of food normalized to 100 g
body weight (Figure 6-4). Leptin treatment decreased food intake in rats fed normal
chow on days 2-6 and 10 (panel A); however, food intake in the leptin-treated group
equaled that in the PBS group by the end of the 2 week implantation period. Leptin had
virtually no effect in rats fed the high-fat (panel B) and had little effect on rats fed the
low-fat diet (panel C).
Effects of Diet and Leptin on Body Weight
The change in body weight was calculated by subtracting the daily body weight from
body weight on the day of pump implantation (prior to surgery) for individual animals,
then an average was taken. Change in body weight was used instead of raw body weight
data due to the differences in starting body weights of the animals within the treatment
groups. In rats fed normal chow, the expected leptin-induced decrease in body weight
was observed beginning on day 3 of leptin administration and continuing for the duration
of the study (Figure 6-5). Leptin treatment in rats fed a high-fat diet, however, attenuated
body weight only on days 10 and 11. In animals receiving the low-fat diet, body weight
was significantly lower in controls starting on day 4 of leptin treatment and continuing
through day 14.
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Effects of Diet and Leptin on IGF-1 Values
In rats fed normal chow, IGF-1 was attenuated by leptin treatment (Figure 6-6) whereas
leptin had no effect on IGF-1 in rats fed the low-fat diet. In rats fed the high-fat diet,
IGF-1 was decreased in both the leptin-treated and PBS-treated groups.
Effects of Diet and Leptin on Organ Weights
Weights of organs were normalized per 100 grams body weight for each individual rat.
There was no difference in weights of brains, pituitaries, hearts, testes, kidneys, livers, or
adrenals in any groups of animals at any time point (Table 6-1).
Effects of Diet and Leptin on Body Temperature
There were no significant differences in body temperature in any groups of rats at any
time point (Table 6-2).
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Figure 6-1: Absolute Food Intake Reported in Various Diet Parameters
Food intake reported as grams of food in rats fed a high-fat diet, a low-fat diet, and normal rat chow (panel
A). Food intake reported as kilocalories in rats fed a high-fat diet, a low-fat diet, and normal rat chow
(panel B). Food intake reported as grams of fat in rats fed a high-fat diet, a low-fat diet, and normal rat
chow (panel C).
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Figure 6-2: Leptin Levels Indicate Proper Pump Activity
In rats fed normal chow serum leptin levels were significantly elevated (p<0.05) at week 2 of leptin
treatment (n=4) (2-way repeated measures ANOVA, panel A). In rats fed the high-fat diet serum leptin
levels were significantly elevated (p<0.05) at weeks 1 and 2 in response to leptin treatment (n=4) (2-way
repeated measures ANOVA, panel B). In rats fed the low-fat diet serum leptin levels were significantly
elevated (p<0.05) at week 2 of leptin treatment (n=4) (2-way repeated measures ANOVA, panel C).
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Figure 6-3: The Effects of Leptin on Food Intake in Diets of Varying Calorie and Fat Contents
In rats fed normal chow food intake was significantly reduced (p<0.05, 2-way repeated measures ANOVA)
in leptin-treated rats (n=4) on days 2-11 and 14 compared to control (n=3, panel A). In rats fed t he high-fat
diet food intake was significantly elevated (p<0.05, 2-way repeated measures ANOVA) in leptin-treated
(n=4) rats on days 7, 12, and 13 compared to control (n=4, panel B). In rats fed the low-fat diet there was
no effect of leptin treatment (n=4) compared to control (p<0.05, 2-way repeated measures ANOVA, panel
C).
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Figure 6-4: The Effects of Leptin on Food Intake - Normalized to 100 grams Body Weight
In rats fed normal chow food intake was significantly reduced (p<0.05, 2-way repeated measures ANOVA)
in leptin-treated rats (n=4) on days 2-6 and 10 compared to control (n=3, panel A). In rats fed the high-fat
diet food intake was significantly reduced (p<0.05, 2-way repeated measures ANOVA) in leptin-treated
(n=4) rats on day 4 compared to control (n=4, panel B). In rats fed the low-fat diet food intake was
significantly reduced (p<0.05, 2-way repeated measures ANOVA) in leptin-treated (n=4) rats on days 2, 3,
and 11 compared to control (n=3, panel C).
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Figure 6-5: The Effects of Leptin on Body Weight in Diets of Varying Calorie and Fat Contents
In rats fed normal chow PBS-treated rats (n=3) gained weight during the 14-day period; leptin-treated rats
(n=4) did not. The results were significantly different (p<0.05) beginning at day 3 of treatment of
continuing for the duration of the study (2-way repeated measures ANOVA, panel A). In rats fed the highfat diet both leptin-treated (n=4) and PBS-treated groups (n=4) gained weight. Body weights were
significantly different (p<0.05) at days 10 and 11 (2-way repeated measures ANOVA, panel B). In rats fed
the low-fat diet both groups gained weight; leptin-treated rats (n=4) gained significantly less (p<0.05) than
PBS-treated rats (n=3) beginning at day 4 and lasting for the duration of the study (2-way repeated
measures ANOVA, panel C).
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Figure 6-6: IGF-1 Values
Leptin treatment (n=3) significantly reduced IGF-1 in normal chow fed rats compared to PBS treatment
(n=3) in the same diet group (p<0.05, 2-way ANOVA). Leptin treatment in the low-fat group (n=3) did not
affect IGF-1 compared to PBS (n=2). In rats fed the high-fat diet, IGF-1 was attenuated in both the leptin(n=4) and PBS-treated (n=3) groups (p<0.05, 2-way ANOVA) compared to the other diets.
Table 6-1: Organ Weights
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Table 6-2: Body Temperature
Discussion
Food intake in rats fed the different diets, before leptin treatment, is reported. The
normal chow rats ingested more grams of food than the other two groups for a majority of
the study, but there were no differences in the intake of kcal among the three groups. As
expected, the high-fat fed rats ingested the highest amount of fat while the low-fat rats
ingested the lowest. These results resemble those seen in Chapter 3 in which the same
diets were utilized.
As was seen in Chapter 5, serum leptin levels increased over time in rats
implanted with leptin-filled osmotic pumps indicating proper pump activity. In the
normal chow and low-fat groups, the elevation of leptin was significant at week 2;
although significance was not reached by week 1 there was an upward trend. In the highfat group, leptin was significantly elevated at weeks 1 and 2. Serum leptin levels were
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not altered in PBS-implanted rats of any diet group over time. It would be expected that
the serum leptin in the PBS-implanted high-fat diet groups would be elevated, however,
beginning body weights were not the same prior to initiation of feeding. As seen in
Chapter 5, steady state serum leptin concentrations were never achieved.
In rats fed normal chow, both food intake and body weight were reduced
beginning on days 2 and 3 of leptin administration, respectively. This effect was
expected and has been well documented in other studies using leptin-filled osmotic
pumps [Pelleymounter et al. 1995; Dawson et al. 1997]. By the end of the two-week
implantation period, however, leptin lost its effects on food intake in the normal chow
rats, indicating that leptin resistance was developing.
When analyzed as grams of food ingested, leptin treatment enhanced food intake
in rats fed the high-fat diet on days 7, 12, and 13. In addition, leptin had no effect on
food intake in the low-fat group. When normalized to grams of food ingested per 100
grams body weight, however, these differential effects of leptin disappear. Leptin
decreases food intake, as expected, in rats fed normal chow; however leptin is much less
effective in rats fed the low-fat diet. In addition, leptin is virtually ineffective in rats fed
the high-fat diet. Taken together, the results of leptin treatment on food intake in rats fed
the different diets suggest that diets higher in calories (both the low-fat and the high-fat
diets) inhibit leptin's effectiveness on this behavior.
Body weight information was reported as change in body weight due to
inconsistencies in weight at the time of implant. Leptin significantly attenuated gain in
body weight in rats fed the low-fat diet for the greater part of the study. Recall that in
normal chow rats, leptin acted to decrease body weight. In low-fat rats, leptin did not
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decrease body weight, but prevented weight gain equivalent to that of the control rats fed
the low-fat diet. Leptin had virtually no effect on body weight in rats fed the high-fat
diet; body weight was attenuated in response to leptin, but only on days 10 and 11.
Leptin regulated body weight more effectively in animals fed a low-fat diet than in
animals fed a high-fat diet, but not as well as in rats fed normal chow. In the case of
body weight, these results suggest diets higher in fat inhibit leptin's ability to curtail body
weight.
It may initially seem illogical that leptin can inhibit gain in body weight when
there is no effect on food intake, as is seen in the low-fat diet group. However, factors
other than those involved with feeding are implicated in the control of body weight. The
rate of metabolism, for example, makes a considerable difference in body composition
among individuals, and leptin has been shown to increase metabolism [Zhang et al. 1994;
Campfield et al. 1995; Halaas et al. 1995; Pelleymounter et al. 1995; Halaas et al. 1997].
Also, there are direct effects of leptin to reduce the size of fat pads [Bai et al. 1996; Qian
et al. 1998]. In the present study, there was no significant increase in body temperature
in leptin-treated rats that would indicate an increased metabolism; however, since neither
VO2max nor activity was measured, an increase in metabolism cannot be ruled out. The
only conclusions that can be made regarding the metabolic status of the rats is related to
IGF-1 levels. However, it was seen in Chapter 5 that the GH-regulating and metabolic
effects of leptin are differentially regulated.
In rats fed normal chow, leptin treatment decreased IGF-1 values, indicating that
the stimulating effects of leptin on GH were reversed. Perhaps the mechanism by which
resistance to the GH-regulating effects of leptin developed was a downregulation of
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hypothalamic leptin receptor as was seen in the previous chapter. Also recall that the
food intake-regulating effects of leptin were lost in these rats. In rats fed the high-fat
diet, IGF-1 was suppressed in both leptin- and PBS-treated groups compared to either of
the two other diets.
Many investigators agree that leptin’s behavioral effects are attenuated in animals
fed a highly palatable diet [Campfield et al. 1995; Frederich et al. 1995a; Masuzaki et al.
1995; Van Heek et al. 1997; Widdowson et al. 1997]. A study by Frederich’s lab
[Frederich et al. 1995a] produced results similar to those reported in the current study:
mice on a “Western” diet of 21% fat demonstrated elevated leptin levels. In Frederich's
study, there was also an increase in body weight. The authors suggested that the high-fat
diet acted to raise the physiological set-point for body weight. Another possibility exists:
that elevated leptin levels produce leptin resistance causing the response to leptin to be
attenuated. This phenomenon was shown in a study in which mice were fed a diet of
45% fat [Van Heek et al. 1997]. These mice experienced elevated serum leptin levels,
grew obese, and were insensitive to peripherally administered leptin. In contrast to the
resistance theory, Campfield et al. [1995] found that leptin, administered twice daily over
two 5-day periods, continued to be effective at altering food intake and body weight in
mice fed a high-fat diet. These results appear to argue against the development of leptin
resistance, however perhaps resistance would have developed had the study been
continued for a longer period of time or with continuous infusion via osmotic minipumps.
Yet another possibility is that leptin's signal is overridden by the ingestion of a high-fat
and/or high-calorie diet.
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In the current study, we observed an attenuation of leptin's effects on food intake
in animals fed diets high in calories (both low-fat and high-fat) and an attenuation of
leptin's effects on body weight and GH secretion in animals fed a diet high in fat. These
phenomena may be the result of an ability of a high-calorie or high-fat diet to impair
leptin’s actions or to raise the body weight set-point defended by the hypothalamus, as
was previously suggested [Frederich et al. 1995a]. Other possibilities exist in which the
high-calorie or high-fat diet simply overrides or antagonizes leptin’s signal by stimulating
other hypothalamic appetite systems [Frederich et al. 1995a] or downregulates leptin
receptors in the blood-brain barrier or hypothalamus.
Numerous "feeding peptides" are nutrient-specific. For example, neuropeptide Y
(NPY) is well-known for its orexigenic effects, especially regarding the intake of
carbohydrates [Stanley et al. 1985; Morley et al. 1987; Bray 1992a; Jhanwar-Uniyal et al.
1993; Wang et al. 1998b] and galanin specifically stimulates the ingestion of fat [Tempel
et al. 1988; Bray 1992a]. Leptin has been shown to inhibit both NPY [Stephens et al.
1995; Schwartz et al. 1996c] and galanin [Sahu 1998] and as such, cannot be said to
negatively regulate any specific type of nutrient.
As a circulating factor informing the brain of the body’s energy stores, leptin
plays an important role in the maintenance of body weight homeostasis. When the brain
fails to recognize leptin, body weight homeostasis is impaired and obesity often develops.
Obese animals and humans have elevated serum leptin levels and are insensitive to the
effects of leptin [Frederich et al. 1995a; Maffei et al. 1995b; Halaas et al. 1997]. This socalled leptin resistance may be the result of one or more circumstances: inability of leptin
to cross the blood-brain barrier [Banks et al. 1996; Caro et al. 1996a; Schwartz et al.
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1996b; Van Heek et al. 1997], defective hypothalamic leptin receptors [Considine et al.
1996a ; Dawson et al. 1997], or impaired post-receptor signaling. The addition to this list
of the ability of a high-fat or high-calorie diet to impair leptin’s actions or to alter body
weight homeostasis regulated by the hypothalamus makes this an important area of study,
especially in today's society where most diets are high in fat and where obesity is
prevalent.
In summary, leptin initially suppressed food intake and body weight in animals
fed normal rat chow; but lost the effects on food intake and IGF-1 by week 2. Leptin
inhibited body weight gain without altering food intake in rats fed a low-fat, high-calorie
diet. In rats fed a high-fat, high-calorie diet, leptin had virtually no effect on food intake
or body weight and IGF-1 was attenuated in both leptin- and PBS-treated subgroups in
these rats. We conclude that leptin’s behavioral, metabolic, and GH-regulating effects
are inhibited by the elevated intake of calories and fat indicating that animals that ingest
such diets lose sensitivity to leptin. The results of this study further support the second
hypothesis of this dissertation.
CHAPTER 7
GENERAL DISCUSSION
In the mammalian endocrine system, a variety of hormones and their hormonal
and metabolic regulators are intricately intertwined, both centrally and in the periphery.
Growth hormone (GH) and leptin are two such hormones. The synthesis of GH in and
the release of GH from the anterior pituitary are stimulated and inhibited by two
hypothalamic neuropeptides, GH-releasing hormone (GHRH) and somatotropin-release
inhibiting hormone (SRIH), respectively. GH has many roles in the body, including the
regulation of energy balance. Leptin, on the other hand, is secreted by adipocytes in
proportion to fat mass. Leptin travels through the bloodstream to the brain where it
informs the hypothalamus of the body's energy stores. The hypothalamus then regulates
food intake and metabolic rate to maintain body weight homeostasis. Both GH and leptin
are modulated by metabolic factors such as feeding, fasting, and obesity. GH and leptin
also act to regulate each other, directly and/or indirectly.
A natural feedback loop involving leptin and GH seems evident. In a normal
physiological situation, the following occurs in the short-term feedback loop: leptin levels
fall, food-seeking behavior is initiated, feeding occurs, leptin levels rise, and feeding is
terminated. In this manner, leptin regulates food intake. In a longer-term feedback loop,
if the effectiveness of leptin to regulate appetite is lost such as can occur in the presence
of a diet high in calories and/or fat, animals may overeat, undergo hyperplasia and/or
hypertrophy of fat cells, and consequently experience elevated levels of leptin. This
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126
elevation in leptin may be sensed at the level of the hypothalamus and/or at the level of
the pituitary to enhance GH secretion. GH stimulates lipolysis, which would then result
in decreased or normalized leptin production. If the loss of effectiveness of leptin on
food intake occurs chronically, leptin resistance develops and the regulation of GH by
leptin may be lost. One of the hypotheses of this dissertation was that in the lean, nonleptin-resistant animal, circulating leptin stimulates the release of GH by acting indirectly
at the level of the hypothalamus and/or directly at the level of the anterior pituitary. The
studies completed in Chapters 3 and 4 support this hypothesis.
In the case presented in Chapter 3, rats were given a high-fat or low-fat diet for 1
month. As expected, rats fed the high-fat diet gained more weight and secreted more
leptin than controls. We consequently saw the predicted elevation in GH secretion,
verified by elevated plasma IGF-1. There was no downregulation of leptin receptor
mRNA in the hypothalamus, possibly indicating that no resistance had developed. The
high fat content of the diet, however, prevent GH's lipolytic actions on body fat and
normalization of leptin levels. This apparent loss of regulation of leptin by GH may have
eventually resulted in the development of leptin resistance had the study been carried out
longer. In fact, this development of resistance in animals fed the same diets over a longer
period of time was seen in Chapter 6, and will be discussed in further detail later in this
chapter.
In Chapter 4, we sought to determine if leptin could produce its effects on GH
directly at the level of the pituitary. A rat pituitary cell line was used in culture to
eliminate indirect (hypothalamic) influence on GH production and secretion. We showed
that leptin receptor mRNA is present in these cells and that leptin treatment significantly
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increased GH secretion at 8 hours under appropriate conditions, in support of the original
hypothesis. The effects of leptin on GH secretion was not significant at 24 hours,
indicating that leptin lost some of its effectiveness over time. With continuous leptin
treatment, leptin receptors on GH1 cells may be downregulated, resulting in a resistance
to leptin.
Leptin resistance had not developed in the studies completed in either Chapter 3
or 4, but in each study there was potential for the development of resistance. In the
pathophysiological situation of leptin resistance, the hypothalamus loses the ability to
detect circulating leptin and therefore continually suppresses GH secretion, as is seen in
obesity. Leptin resistance can develop over time with a malfunction in distribution,
delivery, metabolism, and/or elimination of leptin. It may also occur if leptin
progressively loses its ability to cross the blood-brain barrier, perhaps with a downregulation of the transporter receptors (Ob-Ra) [Caro et al. 1996a]. Additionally, leptin
resistance may occur with downregulation of the long-form of the receptor (Ob-Rb) or
disruption of signal transduction. More than likely, there are multiple pathways involved
in the development of leptin resistance.
The second hypothesis of the dissertation is that in the leptin resistant state, leptin
fails to stimulate GH. In obesity, GH is severely attenuated, and leptin resistance may be
the one of the mechanisms by which this occurs. To support the second hypothesis,
Chapter 5 analyzed leptin resistance as a result of hyperleptinemia due to exogenous
leptin administration. Chapter 6 incorporated the special diets from Chapter 3 and the
induced hyperleptinemia from Chapter 5 to further examine leptin resistance.
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At least one mechanism by which the leptin resistance seen in Chapter 5 occurred
was by a downregulation of hypothalamic leptin receptor mRNA and protein expression.
It is a well-known pharmacological phenomenon that elevated levels of agonist, in this
case circulating leptin, result in downregulation of receptor to prevent excessive activity.
When this occurrence is chronic, there develops a state of resistance to the agonist.
Leptin was infused continuously for 4 weeks via osmotic minipumps at one of
two doses (0.1 mg/kg/day or 0.5 mg/kg/day). The rats in each treatment group became
resistant to the long-term appetite regulating effects of leptin. In addition, both treatment
groups were resistant to the leptin challenge at week 3. Interestingly, the control rats in
the PBS group ate significantly more than the control rats in either leptin treatment group.
These results suggest that, although resistance to the appetite-regulating effects of leptin
was developing in the animals treated chronically, the long-term leptin treatment resulted
in a ceiling effect where the rats could not eat as much as untreated rats. In addition, the
leptin-treated animals were eating the same amount as the untreated animals at this time
point.
In contrast to the resistance to leptin on food intake, the animals in Chapter 5 did
not become resistant to the body weight regulating effects of leptin. Our body weight and
food intake results correspond to those from a study in which mice, after receiving daily
leptin injections, experienced a normalization of the initially reduced food intake
[Pelleymounter et al. 1995]. In that study, the reduction in body weight was never
normalized.
In addition, neither the high-dose nor the low-dose leptin-treated animals
developed resistance to metabolic action of leptin as measured by serum insulin, glucose,
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and triglyceride levels. These results indicate that leptin retained its ability to affect
metabolic parameters even though its ability to alter food intake was lost. It was
previously demonstrated in monosodium glutamate (MSG)-treated rats a dichotomy in
leptin’s actions opposite to that seen in the current study. MSG-treated rats, with damage
to the arcuate nucleus, retained sensitivity to the anorectic actions of leptin, but were
resistant to its metabolic actions [Dawson et al. 1997]. The results of these two studies
suggest that the food intake and metabolic effects of leptin are independently regulated.
Chapter 6 combined the effects of the special diets and exogenously induced
hyperleptinemia to investigate resistance. In Chapter 3, the diets were only fed for 1
month and resistance had not developed so in Chapter 6 the diets were fed for nearly 2
months before experimentation. Osmotic minipumps were implanted and PBS or leptin
(0.25 mg/kg/day) was infused continuously for 2 weeks. Leptin levels were elevated in
rats implanted with osmotic minipumps and also in rats fed the high-fat diet. In animals
fed normal rat chow, leptin initially suppressed food intake and body weight, as would be
expected. However, toward the end of the second week, food intake in the leptin-treated
group was the same as in the PBS group. In addition, IGF-1 was attenuated, indicating
that resistance to the food intake and GH-regulating effects of leptin had developed.
In rats fed the low-fat diet, leptin inhibited body weight gain without altering food
intake. Recall that the low-fat diet was high in calories. Perhaps the caloric content of
the diet was responsible for the diet-induced resistance to the anorectic effects of leptin.
There were no changes in IGF-1 in the low-fat rats. In rats fed the high-fat diet, leptin
had virtually no effect on either body weight or food intake. IGF-1 was suppressed in
both the leptin- and PBS-treated rats fed the high-fat diet. These results suggest that the
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fat content of the diet was responsible for the apparent resistance of the GH-regulating
effects of leptin. No metabolic indices were measured, however, so no statements
regarding rate of metabolism would be substantiated.
The effects of high-fat and/or high-calorie diets on leptin resistance have been
observed previously [Campfield et al. 1995; Frederich et al. 1995a; Masuzaki et al. 1995;
Van Heek et al. 1997; Widdowson et al. 1997]. The ability of a high-calorie or high-fat
diet to impair leptin’s actions or to heighten the body weight set point preordained by the
hypothalamus was previously suggested [Frederich et al. 1995a]. Other possibilities exist
in which the high-calorie or high-fat diet simply overrides or antagonizes leptin’s signal
by stimulating other hypothalamic appetite systems [Frederich et al. 1995a] or
downregulates leptin receptors in the blood-brain barrier or hypothalamus.
It is interesting to consider an opposing hypothesis. In this dissertation, it is
hypothesized that in the normal physiological condition leptin stimulates GH and in the
obese, leptin resistant condition leptin fails to stimulate GH. What if leptin were to
inhibit GH in the physiological condition? Consider the case of a hibernating animal.
During the warmer season, the animal must increase fat stores from which to live during
hibernation. If the fat stores are increasing, leptin levels will also be increasing. It would
be against the best survival mechanisms if GH were being stimulated and reducing fat
stores in these animals. It would seem that, in this instance, the normal physiological
function of leptin would be to reduce GH secretion. However, the physiology of a
hibernating animal is unlike the physiology of a nonhibernating animal. Perhaps the
effects of leptin in an animal preparing itself for hibernation oppose those in
nonhibernating animals. As stated previously, as the animal gains weight in preparation
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for hibernation and circulating leptin levels rise, it would be adverse for lipolysis to
occur. Similarly, decrease of food intake and stimulation of metabolism would be
harmful. The roles of leptin in these animals may be to inhibit GH, stimulate food intake,
and reduce metabolic rate. Alternatively, leptin may have different roles at different
times of year, perhaps under some seasonal circannual control. Yet another possibility is
that the threshold at which leptin resistance develops is lowered in these animals.
Consider another environmental phenomenon: starvation or famine. When food
is scarce, animals are thin and circulating leptin levels would be low. Low leptin
increases the drive to search for food and lowers metabolic rate to conserve energy. It
has been shown that normal levels of leptin are required for normal GH secretion [Carro
et al. 1997], hence in starvation GH is not stimulated by leptin. Both the instances of
famine and hibernation can support the hypotheses proposed in this dissertation.
This dissertation could continue in any of several directions. For example,
additional studies on GH1 cells may include time-course and dose-response studies of
leptin treatment. In addition, leptin receptor concentration could be measured under
normal circumstances and after induction of leptin resistance. Furthermore, the same
experiments could be completed in primary cultures of anterior pituitary cells.
Additionally, supplementary diet studies could be completed in which the effects of
various high-macronutrient diets (high-fat vs. high-carbohydrate vs. high-protein) are
tested on leptin action.
Summary
As a circulating factor informing the brain of the body’s energy stores, leptin
plays an important role in the maintenance of body weight homeostasis. When the brain
132
fails to recognize leptin, body weight homeostasis is impaired and obesity often develops.
Obese animals and humans have elevated serum leptin levels and are insensitive to some
of the effects of leptin [Frederich et al. 1995a; Maffei et al. 1995b; Halaas et al. 1997].
This so-called leptin resistance may be the result of one or more circumstances: inability
of leptin to cross the blood-brain barrier [Banks et al. 1996; Caro et al. 1996a; Schwartz
et al. 1996b; Van Heek et al. 1997], defective [Considine et al. 1996a ; Dawson et al.
1997] or downregulated hypothalamic leptin receptors, or impaired post-receptor
signaling. The addition to this list of the ability of a high-fat or high-calorie diet to
impair leptin’s actions or to heighten the body weight set point preordained by the
hypothalamus makes this an important area of study, especially in today's society where
most diets are high in fat and where obesity is prevalent.
The feedback loop between leptin and GH helps maintain body weight
homeostasis in the normal animal. We have demonstrated the development of resistance
to the anorectic effects of leptin in conjunction with downregulation of hypothalamic
leptin receptor. We have also shown that leptin’s behavioral and metabolic effects are
inhibited by the elevated intake of calories and fat indicating that animals that ingest such
diets lose sensitivity to leptin. We would like to suggest that when this feedback is
disrupted as can occur with consumption of a high-fat diet, resistance to leptin can
develop and obesity can develop. These results may be clinically important in the
prevention and treatment of obesity and must be further explored in basic research. A
possible model of leptin regulation of GH under normal and pathological conditions is
given in Table 7-1.
133
Table 7-1: Model of GH Regulation by Leptin
Physiological
Potential Mechanisms
↓ leptin ↓GH
Insufficient leptin to stimulate GH
↑ leptin ↑ GH
Indirect action via hypothalamic GHRH, SRIH, NPY
Direct action on anterior pituitary somatotropes
Pathophysiological
Potential Mechanisms
↑↑ leptin cannot ↑GH
Leptin resistance
Overriding of leptin action by high-fat and/or high-calorie diet
High-fat and/or high-calorie diet elevation of hypothalamic
body weight set-point
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BIOGRAPHICAL SKETCH
Robin Leigh Picking was born in Crestline, Ohio, on February 20, 1968 to
Thomas R. and Nicolle A. Picking and older brother, T. Reed Picking (age 4). When
Robin was 11, the family moved to Clearwater, Florida, where she attended John F.
Kennedy Middle School and Clearwater High School. Like any teenager, Robin was
anxious to leave the nest and go far from home, so she enrolled in Newberry College in
South Carolina.
The plan to get away backfired, however, when Robin met her future husband,
George Martin, during her first summer break back in Clearwater. Now anxious to return
to Clearwater to be with George, Robin worked hard and finished college one semester
early, graduating with a Bachelor of Science in Biology. In Newberry, Robin enjoyed
some of the best years of her life and made unbreakable friendships, but she looked
forward to living again in Clearwater, not only with George, but also near her parents and
brother.
Back in Florida, Robin worked at a fish hatchery for 7 months and at Pinellas
County Utilities for almost 5 years, during which time she married George, became a
step-mom to Joshua, and bought two cars and house. All along, Robin knew her
education was not complete, so eventually she quit her job and moved to Gainesville to
begin graduate school in the Department of Pharmacodynamics, throwing herself, her
husband, and her step-son into insurancelessness and disrupting all of their lives. George
stayed in Clearwater to be near Joshua, and Robin commuted home on weekends to be
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with her family. Eventually, George also moved to Gainesville, and he and Robin
commuted to Clearwater together to visit the boy on a regular basis.
Robin was very lucky to have stumbled her way into Pharmacodynamics, where
she learned much and had the opportunity to work with many talented people. As the
grand finale to her doctoral academic endeavor draws near, Robin is eager once again to
return to life in Clearwater, and yet, more than a little saddened, for, as in Newberry, her
times in Gainesville have been unforgettable and her friendships developed there
invaluable. She is greatly anticipating the next step in her career, however, and looks
forward to a future in science.
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