The Pituitary (Pituitary (Melmed))

The Pituitary

Notice: The indications and dosages of all drugs in this book have been recommended in the medical literature and conform to the practices of the general community. The medications described do not necessarily have specific approval by the Food and Drug Administration for use in

the diseases and dosages for which they are recommended. The package insert for each drug should be consulted for use and dosage as approved by the FDA. Because standards for usage change, it is advisable to keep abreast of revised recommendations, particularly those concerning new drugs.

The Pituitary EDITED BY

SHLOMO MELMED, Senior Vice-President, Academic Affairs Cedars-Sinai Medical Center Professor and Director, Burns & Allen Research Institute UCLA School of Medicine Los Angeles, California

SECOND EDITION

Blackwell Science

MD

© 2002 by Blackwell Science, Inc. Editorial Offices: Commerce Place, 350 Main Street, Malden, Massachusetts 02148, USA Osney Mead, Oxford OX2 0EL, England 25 John Street, London WC1N 2BS, England 23 Ainslie Place, Edinburgh EH3 6AJ, Scotland 54 University Street, Carlton, Victoria 3053, Australia Other Editorial Offices: Blackwell Wissenschafts-Verlag GmbH, Kurfürstendamm 57, 10707 Berlin, Germany Blackwell Science KK, MG Kodenmacho Building, 7-10 Kodenmacho Nihombashi, Chuo-ku, Tokyo 104, Japan Iowa State University Press, A Blackwell Science Company, 2121 S. State Avenue, Ames, Iowa 50014-8300, USA

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Contents

Contributors vii Preface

Section 3. PITUITARY TUMORS

x

10.

Pituitary Surgery 405 Rudolf Fahlbusch, Michael Buchfelder and Panos Nomikos

11.

Acromegaly 419 Shlomo Melmed

12.

Prolactinoma 455 Mark E. Molitch

13.

Cushing’s Disease 496 Xavier Bertagna, Marie-Charles Raux-Demay, Brigitte Guilhaume, François Girard and Jean-Pierre Luton

14.

Thyrotropin-secreting Pituitary Tumors 561 Yona Greenman and Shlomo Melmed

15.

Gonadotroph Adenomas Peter J. Snyder

16.

Nonpituitary Tumors of the Sellar Region 592 Steffen Albrecht, Juan M. Bilbao and Kalman Kovacs

Section 1. HYPOTHALAMIC–PITUITARY FUNCTION 1.

2.

Functional Anatomy of the Hypothalamic Pituitary Axis 3 Sylvia L. Asa, Kalman Kovacs and Shlomo Melmed Adrenocorticotropin 45 Mark A. Herman and Joseph A. Majzoub

3.

Growth Hormone 79 Vivien S. Herman-Bonert and Shlomo Melmed

4.

Prolactin 119 Mark E. Molitch

5.

Thyroid-stimulating Hormone 172 Virginia D. Sarapura, Mary H. Samuels and E. Chester Ridgway

6.

Follicle-stimulating Hormone and Luteinizing Hormone 216 Shalender Bhasin, Charles E. Fisher and Ronald S. Swerdloff

7.

The Posterior Pituitary Daniel G. Bichet

279

Section 2. HYPOTHALAMIC–PITUITARY DYSFUNCTION 8.

The Hypothalamus 317 Glenn D. Braunstein

9.

Anterior Pituitary Failure Charles F. Abboud

349

575

Section 4. PITUITARY DISEASE IN SYSTEMIC DISORDERS 17.

Pituitary Function in Systemic Disorders 613 Harold E. Carlson

18.

The Pituitary Gland in Pregnancy and the Puerperium 628 Harold E. Carlson

19.

Drugs and Pituitary Function Harold E. Carlson

642 v

vi

Contents

Section 5.

DIAGNOSTIC PROCEDURES

20.

Pituitary Imaging 663 Barry D. Pressman

21.

Neuro-ophthalmologic Evaluation of Pituitary Disorders 687 Anthony C. Arnold

22. Index

Evaluation of Normal Pituitary Function 709 Gillian L. Booth, Afshan Zahedi and Shereen Ezzat 727

Contributors

Charles F. Abboud, MD David Eisenberg Professor of Medicine Consultant, Internal Medicine and Endocrinolgy Mayo Clinic, Mayo Medical School Rochester, MN Steffen Albrecht, MD FRCP(C) Department of Pathology Sir Mortimer B. Davis Jewish General Hospital Montreal, Canada Anthony C. Arnold, MD Associate Professor of Ophthalmology and Chief, Neuro-Ophthalmology Division Jules Stein Eye Institute, UCLA Department of Ophthalmology Los Angles, CA Sylvia L. Asa, MD, PhD Associate Professor of Pathology Mount Sinai Hospital University of Toronto Toronto, Canada Xavier Bertagna, MD Professor of Endocrinology Clinique des Maladies Endocriniennes et Metaboliques Hôpital Cochin Paris, France Shalender Bhasin, MD Chief, Division of Endocrinology, Metabolism, and Molecular Medicine Professor of Medicine Charles R. Drew University School of Medicine and Science Los Angeles, CA

Daniel G. Bichet, MD, FRCP(C) Service de néphrologie et Centre de recherche Hôpital du Sarcé-Coeur de Montréal Department of Medicine Université de Montréal Montréal, Quebec Canada Juan M. Bilbao, MD, FRCP(C) Neuropathologist, Department of Pathology Saint Michael’s Hospital University of Toronto Toronto, Canada Gillian L. Booth, MD Department of Medicine University of Toronto Mount Sinai Hospital Toronto, Canada Glenn D. Braunstein, MD Chairman, Department of Medicine Cedars-Sinai Medical Center Professor of Medicine UCLA School of Medicine Los Angles, CA Michael Buchfelder, MD Associate Professor Department of Neurosurgery University of Erlangen-Nürnberg Erlangen, Germany vii

viii

Contributors

Harold E. Carlson, MD Chief, Endocrinology Section Northport Veterans Affairs Hospital, Northport Professor of Medicine State University of New York at Stony Brook Stony Brook, NY

Kalman Kovacs, MD, PhD, DSc, FRCP(C), FCAP, FRC Path. Professor of Pathology Saint Michael’s Hospital University of Toronto Toronto, Canada

Shereen Ezzat, MD, FRCP(C), FACP Assistant Professor of Medicine University of Toronto Consultant Endocrinologist Wellesley Hospital Toronto, Canada

Jean-Pierre Luton, MD Professor of Endocrinology and Director Clinique des Maladies Endocriniennes et Metaboliques Hôpital Cochin Dean, Cochin Medical School University René Descartes Paris, France

Rudolf Fahlbusch, MD Professor of Neurosurgery Department of Neurosurgery University of Erlangen-Nürnberg Erlangen, Germany Charles Ellis Fisher, MD Division of Endocrinology and Metabolism King Drew Medical Center Los Angeles, CA François Girard, MD Professor of Physiology and Director Laboratoire d’Explorations Fonctionnelles Endocriniennes Hôpital Trousseau Paris, France Yona Greenman, MD Tel-Aviv Sourasky Medical Center Tel-Aviv, Israel Brigitte Guilhaume, MD Clinique des Maladies Endocriniennes et Metaboliques Hôpital Cochin Paris, France Mark A. Herman, MD Fellow in Medicine Department of Internal Medicine University of Texas Southwestern Medical Center Dallas, Texas Vivien S. Herman-Bonert, MD Division of Endocrinology Cedars-Sinai Medical Center Associate Professor of Medicine UCLA School of Medicine Los Angeles, CA

Joseph A. Majzoub, MD Chief, Division of Endocrinology Department of Pediatrics Children’s Hospital Professor of Pediatrics Harvard Medical School Boston, MA Shlomo Melmed, MD Senior Vice-President, Academic Affairs Director, Burns & Allen Research Institute Associate Dean, UCLA School of Medicine Cedars-Sinai Medical Center Los Angeles, CA Mark E. Molitch, MD Professor of Medicine Center for Endocrinology, Metabolism, and Molecular Medicine Northwestern University Medical School Chicago, IL Panos Nomikos, MD Neurosurgical Fellow Department of Neurosurgery University of Erlangen-Nürnberg Erlangen, Germany Barry D. Pressman, MD, FACR Chief, Section of Neuroradiology and Chairman, Department of Imaging Cedars-Sinai Medical Center Los Angles, CA Marie-Charles Raux-Demay, MD Laboratoire d’Explorations Fonctionnelles Endocriniennes Hôpital Trousseau Paris, France

Contributors

Chester E. Ridgway, MD Professor of Medicine and Head, Division of Endocrinology University of Colorado Health Sciences Center Denver, CO Mary H. Samuels, MD Assistant Professor of Medicine Division of Endocrinology Oregon Health Science University Portland, OR Virginia D. Sarapura, MD Assistant Professor of Medicine Department of Medicine, Division of Endocrinology University of Colorado Health Sciences Center Denver, CO

Peter J. Snyder, MD Professor of Medicine University of Pennsylvania School of Medicine Philadelphia, PA Ronald S. Swerdloff, MD Professor of Medicine and Chief, Division of Endocrinology Department of Medicine Harbor/UCLA Medical Center UCLA School of Medicine Torrance, CA Afshan Zahedi, MD Department of Medicine University of Toronto Mount Sinai Hospital Toronto, Canada

ix

Preface

The second edition of The Pituitary follows the successful initial 1995 debut of this comprehensive text devoted to understanding pathogenesis and treatment of pituitary disorders. The new edition is extensively revised to reflect the wealth of novel information derived from advances in molecular biology, biochemistry and therapeutics as they apply to the pituitary gland. Notably, a new chapter devoted to pituitary surgery has been added to complement a comprehensive overview of management options for patients harboring pituitary tumors. The wide spectrum of clinical disorders emanating from disordered function of the “master gland” are described in detail by experts in the field. Furthermore, fundamental mechanisms underlying disease pathogenesis are presented to provide the reader with an in-depth understanding of mechanisms underlying pituitary hormone secretion and action.

Thus the volume continues to reflect the cogent blend of basic science and clinical medicine which was the successful hallmark of the first edition. I am especially indebted to my erudite colleagues for their scholarly contributions and dedicated efforts in compiling this body of knowledge for physicians and scientists dedicated to caring for patients with pituitary disorders. Our desire is to provide medical students, clinical and basic endocrinology trainees, endocrinologists, internists pediatricians, gynecologists and neurosurgeons with a comprehensive single text devoted to the science and art of pituitary medicine. Shlomo Melmed, MD Los Angeles 2002

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Hypothalamic–Pituitary Function

C h a p t e r

1 Functional Anatomy of the Hypothalamic Pituitary Axis Sylvia L. Asa Kalman Kovacs Shlomo Melmed

INTRODUCTION

Historical Review Descartes was the first to recognize that the brain was an organ integrating the functions of the mind and body [1]. The association was reinforced by Zander [2] who noted the connection between the adrenals and the brain, referring to observations on the absence of the adrenal cortex in anencephaly as recorded by Morgagni in 1733, Soemmering in 1792, and Meckel in 1802. It was only in 1849 that direct evidence of a role for the hypothalamus in endocrinology was provided by Claude Bernard [3] when he demonstrated that injury to the floor of the fourth ventricle, the “piqûre diabetique,’ caused polyuria and glucosuria; subsequently, numerous studies in the late 19th century and early 20th century confirmed that the hypothalamic–posterior pituitary system was the site of production of a major osmoregulatory substance. Galen (129–201 ad) had described the pituitary as the site of drainage of phlegm from the brain to the nasopharynx. Soon after the description of acromegaly by Pierre Marie in 1886 [4], the association of acromegaly with pituitary tumor was noted by Minkowski in 1887 [5], and the recognition of endocrine functions of the pituitary followed rapidly thereafter, with major contributions by Cushing [6] and Simmonds [7]. The early part of the 20th century saw the identification, isolation and characterization of the hormones of the anterior pituitary. Their regulation by the hypothalamus was the subject of a landmark monograph by Harris in 1948 [8]. The concept that neurons could release secretory materials into the bloodstream, proposed by Du Bois Reymond

in 1877 [9] and Schiefferdecker in 1905 [10], was revolutionary. The anatomic basis of a hypothalamic–pituitary vascular connection was already known to exist in the form of a system of capillaries, which were identified joining the medial basal hypothalamus to the anterior pituitary by Lieutaud in 1742 [11] and von Luschka in 1860 [12] and given the name “portal plexus” by Popa and Fielding in 1930 [13]. These vessels were thought at first to carry blood from the pituitary upwards to the hypothalamus; however, subsequent studies confirmed that they provided the main blood supply to the adenohypophysis. The concept of neurosecretion was suggested in the 1940s by Scharrer and Scharrer [14] and Bargmann and Scharrer [15], who demonstrated that peptide synthesized by neurons in the supraoptic and paraventricular nuclei of the hypothalamus passed through nerve fibers, was stored in nerve endings in the posterior pituitary and subsequently was released into the general circulation; the analysis and synthesis of oxytocin and vasopressin in 1954 [16] led to a Nobel Prize for du Vigneaud. In the 1950s and 1960s, evidence for the presence of hypothalamic releasing and inhibiting factors accumulated [17]. The isolation and characterization of the first of these yielded shared Nobel prizes for Guillemin and Schally in 1977 and by the mid 1980s, the isolation and characterization of many had been achieved [18–26].

Methodology Progress in the understanding of hypothalamic and pituitary endocrinology has utilized numerous techniques which have allowed the identification of hormones, their isolation and characterization, their localization, and recognition of their cell of origin. At the beginning of the 20th century, Benda [27], Erdheim [28], Cushing [6], and others [29,30] utilized histologic staining methods for the investigation of the human pituitary. Morphologic studies of the hypothalamus were 3

4

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Hypothalamic–Pituitary Function

dependent on conventional neurohistologic methods to demonstrate neurosecretory material and the silver stains that allowed visualization of neuronal cell processes [31]. The tinctorial properties of cell cytoplasm using hematoxylin–eosin, hematoxylin–phloxine, or hematoxylin– phloxine–saffron methods allowed the distinction of three cell types in the adenohypophysis. Clinical observations, correlated with morphologic changes, led to the classification of acidophils as cells thought to produce growth hormone (GH), basophils as those believed to contain adrenocorticotropin (ACTH) and chromophobes as functionally inactive cells. In the 1930s, Rasmussen [32] quantitated the three cell types in relation to sex, age, and body weight and determined their distributions within the gland. In 1940, Romeis [33] used a modification of the trichrome technique to establish that more than three cell types were identifiable in the adenohypo-physis. Application of the periodic acid–Schiff (PAS) method distinguished the PASpositive corticotrophs, thyrotrophs and gonadotrophs; similar results were obtained with lead hema-toxylin. Various trichrome stains, the aldehyde fuchsin and aldehyde thionin techniques were used to separate basophils [34] and the addition of Herlant’s erythrosin [35] and Brookes’ carmoisine [36] were useful to distinguish lactotrophs from the orange G-positive somatotrophs. Physiologic and biochemical studies provided evidence that six different hormones are produced by the anterior pituitary; however, histologic techniques failed to correlate cell type with clinical hormone production or secretory function. The search for correlations between structure and function led to the application of novel techniques including immunocytology [37,38] and electron microscopy [39,40] to the study of pituitary morphology. Ultrastructural immunocytochemistry [41], greatly improved by the use of the immunogold technique [40,42], and the application of the reverse hemolytic plaque assay [43], tissue culture methods [44] and molecular probes [45], have made pituitary and hypothalamic morphology significant for assessing pituitary function and pathology. HYPOTHALAMUS AND POSTERIOR PITUITARY

Topographic Anatomy The hypothalamus is a phylogenetically primitive structure. Understanding of its role in endocrine regulation is largely derived from animal studies; early data regarding functional localization were based on selective stimulation and ablation experiments. However, there are considerable anatomic and physiologic species variations which make extrapolation to the human difficult. Only the advent of sophisticated neurophysiologic methods and the recent applications of immunocytochemistry, as well as experiments of nature in which careful analyses of the location of pathologic lesions are performed, have allowed structure–function correlations in the human hypothalamus.

The hypothalamus is a poorly defined anatomic region which constitutes less than 1% of brain volume and weighs approximately 5 g [46,47]. The superior border of this region is the hypothalamic sulcus which courses from the interventricular foramen to the cerebral aqueduct, demarcating the hypothalamus from the thalamus (Fig. 1.1). The anterior delineation is roughly defined as a line through the anterior commissure, lamina terminalis, and optic chiasm. Posteriorly, it is bordered on the superior aspect by the midbrain tegmentum and inferiorly by the mamillary bodies. The lateral borders are defined conventionally as the substantia innominata, the internal capsule, the subthalamic nucleus, and the cerebral peduncle. The inferior aspect of the hypothalamus, known as the tuber cinereum, contains the median eminence and gives rise to the infundibulum, the neural stalk and the posterior lobe of the pituitary.

Vascular Supply A significant number of afferent and efferent signals of the hypothalamus are not neural, but rather represent bloodborne information, such as temperature, osmotic pressure, hormones, and glucose levels, thus endorsing the importance of the vascular supply of this area. The blood supply of the hypothalamus is derived from small arterial branches of the circle of Willis and the superior hypophysial artery. It is divided into three parts: (i) an anterior group which arises from the internal carotid artery, the anterior cerebral artery, and the anterior portion of the posterior communicating artery; (ii) an intermediate group from the posterior communicating artery; and (iii) a posterior group from the posterior communicating artery, posterior cerebral, and basilar arteries. Individual hypothalamic nuclei receive blood from more than one small artery. The microvascular pattern of the hypothalamus resembles that of the remainder of the brain with the exception of the magnocellular nuclei, the supraoptic and paraventricular nuclei. These are some of the most richly vascularized structures in the brain, in which the neurons are intimately associated with capillaries. The capillary endothelium is fenestrated and the blood–brain barrier is absent in many areas of the hypothalamus, including the subfornical organ, the organum vasculosum of the lamina terminalis (OVLT), the median eminence, and the neurohypophysis. The arterial supply of the median eminence and posterior pituitary has been extensively studied [48–51] (Fig. 1.2). It is derived from two (or three, in some species) paired arteries that arise from the intracranial portions of the internal carotid arteries. The superior hypophysial arteries branch into a plexus of small arteries that surrounds the upper half of the stalk (the external plexus), giving rise to a mesh of capillaries, and the gomitoli (the internal plexus). The latter are unique vascular structures composed of a central muscular artery surrounded by a spiral of capillaries, the arteriole communicating with the capillaries by way of small

Chapter 1

Functional Anatomy of the Hypothalamic Pituitary Axis

5

FIGURE 1.1. Schematic representation of the human hypothalamus and pituitary, the disposition of hypothalamic nuclei and principal fiber tracts. From Scheithauer [47]

orifices surrounded by muscular sphincters. The gomitoli, which measure 1–2 mm in length and 0.1 mm in width, are found in large numbers in the infundibulum and proximal hypophysial stalk. Flow through these complex structures continues on through the portal vessels to the adenohypophysial capillaries. Although their function is not certain, the unique structure of gomitoli suggests that they may affect the rate of blood flow to the anterior pituitary, thereby influencing the entry of hypothalamic regulatory hormones into the portal circulation. In some individuals, a pair of middle hypophysial arteries also contributes to the trabecular or loral arteries which descend from the superior hypophysial arteries along the external surface of the pituitary stalk. These then divide to contribute to the subcapsular artery and the artery of the fibrous core (which provide a minor contribution to the blood supply of the adenohypophysis) and return upwards along the pituitary stalk as the long stalk arteries. The inferior hypophysial

arteries enter the sella just beneath its diaphragm and supply the pituitary capsule, the neural lobe, and the lower pituitary stalk. They enter the intralobar groove where they divide into ascending and descending branches that join their opposite partners to form an arterial circle about the neural lobe. They also give off a branch to the lower pituitary stalk, the communicating artery, which anastomoses with the trabecular arteries. Notably, the capillaries of the neurohypophysis are fenestrated and lie outside the blood–brain barrier. The earliest studies of the hypophysial portal vasculature assumed that blood flowed from the pituitary upwards to the hypothalamus [13]. Subsequently, it was found that the predominant blood flow is from the hypothalamus to the adenohypophysis, carrying hypothalamic regulatory factors. Recent studies suggest that the direction of blood flow is variable and there may be a component of reverse flow within the neurohypophysis, providing a route by which

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FIGURE 1.2. Schematic representation of the blood supply of the hypothalamus and pituitary. From Scheithauer [47]

neuro- and adenohypophysial hormones gain access to the long portal system, the pars tuberalis of the adenohypophysis, the median eminence, and the cerebrospinal fluid of the third ventricle [50,51].

Nerve Supply A number of afferent neural pathways provide the hypothalamus with input from the forebrain, limbic system, visual cortex, thalamus, and brain stem [47,52]. The median forebrain bundle, a large unmyelinated structure lateral to the fornix, carries information to and from the brain stem and provides important input from the limbic system and olfactory forebrain structures. Olfactory connections are more extensively from the piriform cortex and

amygdaloid nuclear complex via the stria terminalis and ventral amygdalofugal pathway. The hippocampal–hypothalamic tract originates in the hippocampus, a limbic structure, and enters the anterolateral portion of the mamillary bodies. The amygdalo–hypothalamic tract provides entry of emotional data from the amygdaloid nucleus into the hypothalamus. The retino–hypothalamic tract originates in the ganglion cells of the retina and affects the pineal gland, playing a role in the regulation of circadian rhythms. In addition, a small number of cortico–hypothalamic fibers from the posterior orbital gyrus of the frontal lobe is a potential source of input from the neocortex into the hypothalamus.

Chapter 1

The hypothalamus sends efferent projections to the forebrain, brain stem, spinal cord and neurohypophysis. The median forebrain bundle carries fibers to the septal nuclei, integrating visceral and olfactory functions. The dorsal longitudinal fasciculus carries efferents to the midbrain tegmentum where it innervates visceral and sympathetic motor neurons. Fibers emanating from the mamillary body divide into the mamillo–thalamic tract, serving the anterior nucleus of the thalamus and memory function, and the mamillotegmental tract which projects to the midline tegmentum. Efferents from the ventral medial nucleus project to the amygdaloid nucleus via the stria terminalis. The hypothalamo–hypophysial tract consists primarily of nerve fibers from the supraoptic and paraventricular nuclei carrying vasopressin, oxytocin and their neurophysins to the posterior lobe of the pituitary, where the hormones are released into capillaries. The tubero–infundibular tract, originating from neurosecretory neurons which produce hypophysiotropic hormones, projects from several nuclei to the median eminence where the hormones are released into the hypophysial portal vascular system.

Nuclei and Nucleoinfundibular Pathways Within the hypothalamus, distinct clusters of neurons are called “nuclei.” The hypothalamic nuclei are generally divided into four anatomic regions, the preoptic, supraoptic and lateral, tuberal, and mamillary regions. In general, the nuclei are topographically discrete in many species and, although relatively well demarcated in the human fetus, they are poorly defined in the mature human hypothalamus [46]. In a few cases, the existence of some nuclei is inferred entirely from studies of experimental animals. A schematic illustration is provided in Fig. 1.1. Structure–function correlations are difficult because of cellular heterogeneity of many hypothalamic nuclei. Any given hypothalamic hormone is often produced in more than one nucleus and, in many cases, a single nucleus may express more than one hormone. The physiologic roles of many nuclei remain unknown. The most anterior nuclei are the paired medial and lateral nuclei that have been associated with autonomic function, particularly temperature control and olfaction. The suprachiasmatic nucleus is found in the preoptic area dorsal to the optic chiasm and anterior to the supraoptic nucleus. This sexually dimorphic cell group shows a striking decrease in volume and cell number with age [53]. It is located in an area that is essential for gonadotropin release and sexual behavior in lower animals and may play a role in maintaining circadian rhythms. It is thought that the nucleus also plays a role in the sexual differentiation of the brain which, in the absence of male gonadal hormones, remains female but, if exposed to these at a critical stage in development, becomes male [54]. It is also associated with fibers of the supraoptic commissure, and receives afferents from the retina and the lateral geniculate bodies.

Functional Anatomy of the Hypothalamic Pituitary Axis

7

The anterior hypothalamic nucleus is composed of small neurons which mediate parasympathetic effects; it has connections to the insular cortex, substantia innominata and thalamus. The lateral hypothalamic nuclei are poorly defined areas composed of larger neurons which receive fibers from and contribute efferents to the median forebrain bundle. The paraventricular nuclei lie adjacent to the third ventricle and ventromedial to the fornix. They are composed mainly of large “magnocellular” neurons along with the supraoptic nuclei and also contain a number of “parvicellular” neurons. Efferent fibers to the pituitary stalk terminate in the posterior lobe of the pituitary. These nuclei are a major site of oxytocin and vasopressin synthesis. In patients with traumatic or surgical stalk section and in those with long-standing hypopituitarism, there is a marked reduction in the number of magnocellular neurons and stalk nerve fibers [46]; the parvicellular component remains. The supraoptic nuclei are the other paired magnocellular nuclei of the hypothalamus; they have no significant parvicellular component. They overlie the anterior optic tract and posteriorly are divided by the optic tract into anterolateral and posteromedial components, united by a thin stream of cells. Patients with stalk section or long-standing hypopituitarism also have atrophy of these nuclei [55]. The dorsomedial and ventromedial nuclei are involved in autonomic function and emotional behavior. They are situated between the tuber cinereum and paraventricular nuclei. Stimulation of the dorsomedial and destruction of the ventromedial nuclei produces rage in experimental animals. The ventromedial nuclei are concerned with control of balance between hunger and satiety. Destruction of the ventromedial nucleus results in obesity [56,57]; conversely, destruction of the ventrolateral nucleus, known as the “feeding center,” is associated with anorexia and cachexia [56]. These nuclei show a large number of afferent connections from olfactory and retinal fibers, the reticular formation, and the nucleus of the solitary tract which receives input from the vagus. Afferents from the cortex enter by way of the thalamus. Ventral to the third ventricle and paraventricular nuclei lies the arcuate (infundibular) nucleus which is another important component of the hypophysiotropic region and plays a major role in the modulation of anterior pituitary function. The subventricular nucleus lies on the floor of the third ventricle posterior to the arcuate and anteromedial to the tuberal nuclei. This usually parvicellular nucleus shows marked hypertrophy of the neurons which become magnocellular in postmenopausal women [58,59], in young women suffering from postpartum hypopituitarism with complete gonadal atrophy [60], in hypogonadal men and women, in starvation [61], after hypophysectomy [62], and in late pregnancy. The neurons develop a distinctive nucleolar change considered a manifestation of feedback effect; a similar change is also observed in neurons of the arcuate

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Hypothalamic–Pituitary Function

nucleus [60]. In some of these conditions, the change is thought to reflect lack of estrogens [59,63,64]. The tuberal nuclei consist of irregularly grouped masses of large neurons inferior to the lateral nuclei that give rise to efferent fibers of the hypothalamus. After stalk section, a slight increase in coarsely granular basophilic cytoplasmic material may occur [46]. The posterior hypothalamic nucleus, situated between the third ventricle and the mamillo–thalamic tract, lies superior to the mamillary bodies. When stimulated, it produces sympathetic effects; it may play a role in temperature regulation and its large neurons may be the source of hypothalamic efferents that descend to the reticular formation of the brain stem. The paired mamillary nuclei and a variety of minor nuclei in the supramamillary area, including the nucleus intracalatus, form the most posterior portion of the hypothalamus. These nuclei integrate incoming information from the limbic system and the midbrain tegmentum and send out efferent fibers to the anterior nucleus of the thalamus and to the brain stem.

Embryology The forebrain of the human embryo can be identified at 4 weeks postconception; by 5 weeks it has divided into a central diencephalon and two lateral telencephalic structures. Three swellings appear in the diencephalon which represent the anlages of epithalamus, thalamus, and hypothalamus. The nuclei of the hypothalamus and the supraoptico-hypophysial tract are recognizable at 8 weeks of gestation [65]. The nerve cells originate from proliferating neuroblasts and form several closely associated nuclei in the hypothalamus. At 6 weeks, a thickening of neuroblasts projects caudally to become the median eminence and infundibulum and establishes direct contact with the upward invagination of the roof of the primitive oral cavity, Rathke’s pouch. By 3 months, the rudimentary neural lobe consists of a short wide sac with a thick wall and a lumen lined by columnar ependymal cells [66]. Proliferation of spongioblasts gives rise to the modified neuroglia of the neurohypophysis known as pituicytes while the ependymal cells become dispersed in the neuroglial substance. Downgrowth of unmyelinated axons of neurons of magnocellular nuclei of the hypothalamus reaches the median eminence by 3 months, passes through the stalk and terminates in the neural lobe by 6 months of gestation. Monoamine fluoresence can be detected at 10 weeks in hypothalamic nuclei and at 13 weeks in the median eminence [67]. The supraoptic nucleus matures earlier than the paraventricular nucleus; vasopressin is demonstrated earlier and in higher concentrations than oxytocin in the neural lobe [68]. Arginine vasopressin is identified by radioimmunoassay at 12 weeks of gestation [69]. Intracellular granules have been demonstrated in hypothalamic nuclei at 19 weeks and in the posterior lobe at 23 weeks of gestation. The

neurosecretory material is clearly visible in the supraoptic and paraventricular nuclei as well as the posterior lobe at 23–28 weeks of gestation [70–72]. Ultrastructurally, clusters of small clear vesicles and axon profiles have been identified in a fetus at 7.5 weeks of age. Granular vesicles become apparent in the axon profiles at 8.5 weeks of gestation. Fenestration of capillaries and expansion of perivascular spaces are noted with the appearance of intraaxonal vesicles [73]. Growth hormone-releasing hormone (GHRH) has been identified by immunohistochemistry in the fetal hypothalamus at 18 weeks of gestation [74]. Somatostatin (somatotropin release-inhibiting hormone; SRIH) is identified in fetal hypothalami in a large molecular form at 10 weeks of gestation and there is a gradual increase in both the number of somatostatin-containing neurons and the 14-amino acid form of this peptide with fetal age [75,76]. Dopamine is present in high concentrations in the fetal hypothalamus by 15 weeks of gestation [67]. Corticotropin-releasing hormone (CRH) has been detected in fetal median eminence at 16 weeks of gestation [77]. Thyrotropin-releasing hormone (TRH) and gonadotropin-releasing hormone (GnRH) are the first adenohypophysiotropic substances, being detectable in fetal hypothalami at 4.5 weeks and present in significant amounts by 10 weeks of gestation [78,79]. GnRH is localized by immunocytochemistry at 13 weeks and GnRH-containing nerve terminals are detected on portal vessels at 16 weeks [80]. Sexual dimorphism of GnRH concentration has been documented and correlates with sexual dimorphism of fetal gonadotropic hormones [81].

Functional Anatomy and Control of Anterior Pituitary Function Control of anterior pituitary hormone secretion is under a complex set of central and peripheral neural and chemical signals. The hypothalamus synthesizes and secretes unique releasing and inhibitory hormones that control the anterior pituitary hormones (Fig. 1.3). The polypeptide hormones are secreted into the hypothalamic pituitary portal vasculature, impinge upon specialized anterior pituitary cells and bind to specific surface receptors. The cell signal elicits regulation of pituitary trophic hormone gene transcription, translation, and/or secretion. The resultant secretion of anterior pituitary hormones into the systemic circulation depends upon a carefully controlled dynamic balance of hypothalamic hormone input, feedback regulation from peripheral target hormones, nonhormonal neurotransmitter agents, and paracrine or autocrine pituitary growth factors and peptides. These factors all contribute to the amount of each pituitary hormone secreted, as well as to their pattern of secretion. Complex feedback regulation loops interact to regulate the hypothalamic–pituitary axis. The target gland peripheral hormone usually exerts negative feedback regulation directly at the level of pituitary synthesis or secretion of the respective trophic hormone (Fig. 1.4). Pituitary

Chapter 1

Functional Anatomy of the Hypothalamic Pituitary Axis

9

FIGURE 1.3. Amino acid sequences of human hypothalamic hormones. CRH, corticotropin-releasing hormone; GHRH, growth hormone-releasing hormone; GnRH, gonadotropin-releasing hormone; SRIH, somatostatin; TRH, thyrotropin-releasing hormone.

FIGURE 1.4. General control of anterior pituitary hormone release by central and hypothalamic inputs and feedback regulation by peripheral and trophic hormones. ACTH, adrenocorticotropic hormone; CRH, corticotropin-releasing hormone; DA, dopamine; FSH, folliclestimulating hormone; GHRH, growth hormone-releasing hormone; GnRH, gonadotropin-releasing hormone; IGF-I, insulin-like growth factor-I; LH, luteinizing hormone; PRL, prolactin; SRIH, somatostatin; TRH, thyrotropin-releasing hormone; TSH, thyrotropin; VIP, vasoactive intestinal polypeptide.

trophic hormones may feed back in a very short loop to autoregulate their own secretion. They may also participate in short loop feedback regulation of the secretion of their respective hypothalamic releasing or inhibiting hormone. Finally, peripheral hormones may participate indirectly by feedback regulation of hypothalamic releasing and inhibiting hormones.

The central nervous system also influences the control of hypothalamic hormonal secretion by efferent signals emanating from exteroceptive organs including the ears, eyes, and skin, and from interoceptive senses of indices of metabolic and electrolyte homeostasis. The transduction of neuronal inputs to neurochemical signaling by the hypothalamus appears to occur in an oscil-

10

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latory fashion. The temporal nature of pituitary hormone secretion is a manifestation of this rhythmicity resulting in at least two patterns of circadian rhythm including sleeprelated hormone secretory cycles and 24-hour rhythm cycles entrained by a light–dark cycle. The peptidergic neurons of the hypothalamus are not classified into distinct cell types and there is no known correlation between cell structure and function. In fact, neurons of any anatomic area often contain more than one neuropeptide or candidate neurotransmitter. The functional anatomy of the hypothalamus is, therefore, best approached by discussion of each hormone or peptide. GHRH

The presence of the factor responsible for stimulating release of growth hormone (GH) was known for many years, but it was only in 1982 that GHRH was isolated and characterized [25,26]. It is now known to have two forms, a 40- and a 44-amino acid peptide; the active portion of the molecule is the 29-amino acid N-terminus. GHRH binds to the surface of the somatotrophs. The intracellular actions of GHRH are probably mediated by cyclic adenosine monophosphate (cAMP), resulting in stimulation of GH gene transcription, translation, and secretion. Somatostatin attenuates the secretion of GH by the somatotroph. GH stimulates predominantly the liver to produce insulin-like growth factor-I (IGF-I) which acts as a target hormone to induce tissue growth. GH itself also acts directly on the chondrocytes and possesses antiinsulin metabolic activity. Immunocytochemical studies have identified GHRH in neuronal cell bodies of the arcuate nucleus [82,83] (Fig. 1.5). Nerve fibers from this nucleus project mainly to the median eminence, are found in the external zone, and terminate on capillaries of the plexus of the hypothalamohypophysial portal system. In monkey hypothalamus, small groups of immunoreactive neurons are also found in ventral

portions of the ventromedial nucleus. In the rat, scattered cells are also seen in the lateral basal hypothalamus, the medial and lateral portion of the ventromedial nucleus, and the dorsal medial and paraventricular nuclei [84,85]. The primary anatomic localization of GHRH to the arcuate nucleus is in agreement with earlier physiologic studies which indicated that a region including the ventromedial and arcuate nuclei was facilitatory in GH regulation. Extrapituitary effects of GHRH have been reported in the rat. Administration of small doses of the peptide into the cerebral ventricles induces behavioral sedation as well as an increase in food intake [86]. No extra-hypothalamic sites of GHRH synthesis are known to exist in the human under normal circumstances, but endocrine tumors of lung, pancreas, gut, adrenal medulla, and C-cells of thyroid can give rise to GHRH excess and acromegaly [87] (see Chapter 11). Somatostatin (SRIH)

A hypothalamic factor with GH release-inhibiting activity was first described in 1968 [88]. In 1973, Brazeau and colleagues [23] isolated and characterized a 14-amino acid peptide which they called somatostatin, or somatotropin release-inhibiting hormone (SRIH). This peptide was also shown to inhibit thyrotropin (thyroid-stimulating hormone; TSH) secretion and to produce a variety of other effects not related to its action on the pituitary. The hypothalamic distribution of SRIH shows marked species variation [89–92]. In the human hypothalamus, SRIH-containing neurons are concentrated in the anterior portion of the arcuate nucleus [90]. No positivity has been found in the paraventricular nucleus, unlike the distribution observed in rodents where SRIH is found in the paraventricular nucleus in a narrow field extending 1–2 mm from the midline. Nerve fibers staining for SRIH are observed in the medial basal hypothalamus; nerve endings are concentrated in the external zone of the median eminence and the neurovascular zone of the pituitary stalk. Fibers containing SRIH are also localized in the ventromedial, arcuate, suprachiasmatic, and paraventricular nuclei. Somatostatincontaining neurons store the hormone in dense core cytoplasmic vesicles which measure 80–110 nm in diameter and are found in both the cell bodies and the nerve endings [92]. Somatostatin is also found in extrahypothalamic regions of the rat brain, including the preoptic region, central gray mesencephalon, amygdala, olfactory area, and spinal cord. In humans it is detectable in cortex, brain stem, pineal, retina, optic nerve, auditory nerve, and spinal cord. Somatostatin is also found in pancreatic islet cells, gut endocrine cells, thyroid C-cells, salivary gland, and placenta [93–95]. Prolactin-inhibiting Factor

FIGURE 1.5. Neurons containing immunoreactive growth hormone-releasing hormone are localized by immunocytochemistry in the arcuate nucleus. (Magnification ¥64)

Prolactin (PRL) secretion by the adenohypophysis is known to be under tonic inhibition by the hypothalamus [96]. The PRL-inhibiting activity of dopamine is well documented and, currently, it is widely accepted that dopaminergic

Chapter 1

control is the main PRL regulatory function of the hypothalamus. Dopamine is the only nonpeptidergic hypothalamic compound with a well-defined hypophysiotropic function. It exerts primarily inhibitory effects on the lactotroph and inhibits both PRL gene transcription and lactotroph mitotic activity. PRL acts to induce lactation in concert with other peripheral hormones. Although TRH also stimulates lactotroph function, its effect is probably not physiologic. Vasoactive intestinal peptide (VIP) has also been shown to stimulate the production of PRL although its physiologic role has not yet been clarified. Dopamine has been localized to the arcuate and periventricular nuclei of the mediobasal hypothalamus as well as being widespread throughout the brain [97,98]. The medium-sized dopaminergic neurons are scattered among other neurosecretory cells and are further subdivided into the tuberoinfundibular group, with terminals in the median eminence and pituitary stalk, and the tuberohypophysial group, with ends terminating in the neural and intermediate lobes of the pituitary. The dopamine projections to the intermediate and neural lobes of the pituitary originate primarily in the anterior and central portions of the arcuate nucleus, respectively. Dopamine-containing axon terminals are abundant in the zona externa of the median eminence and are characterized by dense core vesicles that measure 15–120 nm in diameter. They do not appear to form true synaptic connections. CRH

Despite being the first hypothalamic regulatory factor postulated, CRH was among the last identified. In 1981, a 41-amino acid peptide was extracted from the ovine hypothalamus, characterized and shown to stimulate specifically the release of ACTH and other pituitary peptides derived from proopiomelanocortin (POMC) [24]. CRH binds to the surface of the corticotroph and elicits transcription of the POMC gene, whose products include ACTH and bendorphin. ACTH stimulates the adrenal gland to synthesize corticosteroids which are critical for essential stress responses and cellular homeostasis. Antibodies against CRH have localized this peptide to neuronal cell bodies, mainly in the parvocellular portion of the paraventricular nucleus but also in a few magnocellular neurons [99–104]. In the rat hypothalamus, scattered cell bodies containing CRH are also detected in the lateral preoptic and hypothalamic areas and in some portions of the dorsal medial nucleus. CRH-immunoreactive fibers are observed in the area of the paraventricular nucleus, the neurovascular zone of the median eminence, the pituitary stalk, and the neurohypophysis. CRH is colocalized with arginine vasopressin in a subpopulation of magnocellular neurons of the paraventricular nucleus [103,104]; it is found together with enkephalin, peptide histidine isoleucine (PHI, a member of the secretin–glucagon family) and neurotensin in parvocellular neurons as well as in a discrete subset of oxytocinergic magnocelluar neurons [104]. Adrenalectomy

Functional Anatomy of the Hypothalamic Pituitary Axis

11

increases CRH immunoreactivity in parvocellular neurons of the paraventricular nucleus, consistent with established negative feedback effects of adrenal steroids on CRH production [104]. Vasopressin can also be found in the majority of CRH-containing parvocellular neurons after adrenalectomy [104]; it has been suggested that this plasticity allows for synergy of stimulation of ACTH secretion by the two peptides [104]. CRH is also widely distributed throughout the brain in the cerebral cortex, limbic system, brain stem, and spinal cord [98–101]. It is also found extracerebrally, particularly in placenta; some investigators have also localized CRH in endocrine cells of the pancreas, gut and lung; in liver, adrenal [105], and testicular Leydig cells [106]. TRH

The first adenohypophysiotropic hypothalamic hormone identified was the tripetide TRH [17,18]. At first demonstrated to release TSH, TRH was rapidly also shown to release PRL. TRH binds to specific receptors on the thyrotroph and stimulates the transcription and secretion of TSH. TSH stimulates the thyroid gland to secrete primarily thyroxine, and triiodothyronine. The major source of triiodothyronine, however, is hepatic deiodination of thyroxine. These hormones act to regulate thermogenesis and protein synthesis. Immunocytochemical studies have localized TRH in the rat brain, in neuronal cell bodies of the preoptic nucleus, the parvocellular portion of the paraventricular nucleus, the perifornical region, dorsomedial nucleus, and basolateral hypothalamus [107,108]. TRH-positive terminals that contain dense core vesicles are found in high concentrations in the median eminence as well as in several of the hypothalamic nuclei, which also contain TRH-positive neuronal cell bodies. The tripetide is widely distributed throughout the brain and spinal cord, suggesting that it may act as a neurotransmitter in other areas [107,108]. Several effects of TRH not mediated by the pituitary gland have been demonstrated, including hyperthermia, behavioral excitation and the ability to reverse or prevent anesthesia from barbiturates and other depressants [109]. TRH has been reported in fetal and neonatal pancreatic islet cells [110] and gut endocrine cells [111], where it is thought to inhibit gastric secretion, gastric motility, and exocrine pancreatic secretion; it has also been localized in several human tumors [112]. GnRH

The second hypothalamic adenohypophysiotropic peptide to be characterized was originally called luteinizing hormone (LH)-releasing hormone (LRH) but was subsequently shown to release both gonadotropins, follicle stimulating hormone (FSH), and LH, by the pituitary gland [20–22]. The gonads are thereby stimulated to produce sex steroids and secondary sex characteristics. GnRH is also thought to stimulate sexual activity in experimental animals of both

12

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Hypothalamic–Pituitary Function

sexes, an effect that appears to be independent of its hypophysiotropic action [113]. GnRH has been localized in the hypothalami of several species [113–115]. In primates, GnRH neurons are located in large areas extending caudally from the preoptic region to the premamillary nucleus [96]. The neurons are bipolar or multipolar cells with multiple dendrites. The highest concentration of these cells is found in the mediobasal hypothalamus (infundibular and premamillary nuclei) and in the preoptic area. These neurons give rise to an infundibular GnRH pathway ending on the capillaries of the pituitary portal plexus as well as to a preopticoterminal tract projecting to the capillaries of the OVLT. In rodents, the majority of neuronal cell bodies are located in the septal preoptic regions and the anterior hypothalamus, and are particularly in the dorsal and medial septal nuclei, the nucleus of the diagonal band of Broca and anterior hypothalamic nuclei, and the OVLT. These neurons are usually elongated with eccentric nuclei and are unipolar. They project to the external zone of the median eminence as well as to the OVLT. Immunoelectron microscopy has documented localization of GnRH in dense core vesicles which measure 75–95 nm in diameter and are found in cell bodies as well as nerve endings [115]. Extrahypothalamic localization of GnRH includes several areas of the limbic system (septum, hippocampus, and olfactory bulb), the breast, and placenta [116]. Vasopressin and Oxytocin

Vasopressin, oxytocin, and their binding proteins, neurophysins I and II, are detected by immunohistochemistry in the perikarya of magnocellular neurons in the supraoptic (Fig. 1.6) and paraventricular (Fig. 1.7) nuclei [117,118]. Fibers emanating from these neurons converge towards the median eminence, pass caudally through its internal zone and end in dilated nerve terminals of the pars nervosa of

the pituitary known as Herring bodies (Fig. 1.8). The nerve fibers, their axonal swellings and the exceedingly large and characteristic swellings which can exceed 20 mm in diameter contain secretory granules of variable electron density and electron-lucent vesicles (Fig. 1.9). The secretory granules originate in the perikarya of the neurons and mature, growing from 60 to 220 nm in diameter, as they travel to the nerve endings [119,120]. It has been clearly shown that vasopressin and oxytocin are produced by different magnocellular neurons in both nuclei. Immunostaining for vasopressin is markedly increased after adrenalectomy in the external zone but not the internal zone of the rat median eminence [121,122]. Vasopressin is colocalized in some neurons with CRH and is additive with that substance in the regulation of ACTH release. Interestingly, it is found in high concentrations in portal blood draining to the anterior pituitary, suggesting that it may play a physiologic role in ACTH release [104]. A subset of oxytocin-containing neurons also may contain CRH. Immunoreactive vasopressin has been demonstrated in various other areas of the central nervous system as well as testis, ovary, uterus, adrenal, and thymus [123]. VIP

This 28-amino acid peptide, first isolated from intestine, has been localized in mammalian brain [124]. Immunoreactive fibers are present in high concentrations in the cortex, hippocampus, dentate gyrus, amygdala, nucleus of the stria terminalis, suprachiasmatic nucleus, and periaqueductal gray matter. Fibers containing immunoreactive VIP and its related PHI are also detected in areas that do not contain cell bodies. In the hypothalamus, VIP has been localized in the ventromedial nucleus, where it may affect appetite [56], in the suprachiasmatic nucleus, where it is thought to play a role in circadian rhythmicity, and in parvocellular neurons of the paraventricular nucleus [125]. It is known to modulate secretion of GH, PRL, and LH at the level of their hypothalamic regulation [126–128] and also stimulates PRL release directly [126]; some investigators have suggested that VIP is the physiologic PRL-releasing factor. After adrenalectomy, the number and intensity of VIP-containing small neurons in the paraventricular nucleus increase, suggesting a possible role for this substance in PRL regulation during stress. Cholecystokinin (CCK)

FIGURE 1.6. Perikarya and axonal processes of magnocellular neurons in the supraoptic nucleus contain strong immunopositivity for vasopressin. (Magnification ¥64)

This hormone, which was first discovered in the gut, was detected in the brain by immunocytochemistry [129–131]. CCK-containing cell bodies are prominent in the cerebral cortex and are also widely distributed throughout the olfactory and limbic systems. In the hypothalamus, they are found in several nuclei, including the magnocellular paraventricular and supraoptic nuclei. The midbrain also contains a striking concentration of CCK-immunoreactive cell bodies in the ventral tegmentum and raphe. This peptide modulates dopaminergic, noradrenergic, and opioid systems. It also inhibits feeding, probably at the level

Chapter 1

Functional Anatomy of the Hypothalamic Pituitary Axis

13

FIGURE 1.7. Adjacent to the third ventricle (V), magnocellular neurons in the paraventricular nucleus demonstrate strong immunoreactivity for vasopressin. (Magnification (a) ¥26; (b) ¥64)

FIGURE 1.8. The posterior lobe of the pituitary contains nerve fibers of the hypothalamohypophysial tract with axonal swellings and globular Herring bodies that stain with aldehyde thionin (a) and contain intense immunopositivity for vasopressin (b). (Magnification (a) ¥102; (b) ¥64)

of the ventromedial hypothalamus, and alters the pain threshold [132].

release, and to enhance GH release in response to GHRH in humans and rats [141], suggesting that hypothalamic galanin may modulate secretion of those hormones.

Galanin

Galanin is a 29-amino acid peptide that was originally isolated from intestine [133]. Immunoreactive galanin has been demonstrated in the central nervous system [134]; in hypothalamus, it colocalizes with dopamine and g-aminobutyric acid (GABA) [135]. It is also present in peripheral neural elements of the respiratory tract, gastrointestinal tract, pituitary, pancreas, urogentital tract, and adrenal medulla of humans and several mammalian species [136–139]. Galaninpositive neurons have been observed in several hypothalamic tumors [139]. Galanin is thought to act as a neurotransmitter although its functional roles have not been fully clarified. It has been shown to inhibit dopamine [140], thereby stimulating PRL

Gastrin

Gastrin is a gut peptide which is known to be produced in stomach and in numerous endocrine tumors of gut and pancreas. Specific antibodies have also localized gastrin in cerebral cortex, hypothalamus, and neurohypophysis [142,143]. Gastrin is known to modulate GH secretion at the hypothalamic level, either by reducing somatostatin release or by enhancing GHRH secretion [144]. Glucagon

Glucagon is expressed in the pancreatic islets and intestine, and has also been reported in other tissues, including thymus, thyroid, and adrenal medulla. A growing body of

14

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Hypothalamic–Pituitary Function

levels of NPY immunoreactivity are found in hypophysial portal blood. This peptide stimulates appetite when injected into the hypothalamus and appears to modulate the secretion of both LH and ACTH [150]. NPY-containing fibers also are known to contact TRH-synthesizing perikarya and neuronal processes in the paraventricular nucleus where it may mediate the neuroendocrine regulation of TRH [151]. Neurotensin

This 13-amino acid gut–brain peptide is found in the highest concentration in the ileum and hypothalamus [152]. In the rat, it is localized in the medial preoptic region, infundibular nucleus, paraventricular nucleus, and lateral hypothalamus with fibers most dense in the paraventricular and periventricular regions and in the median eminence. It has been shown to colocalize in neurons with catecholamines, possibly dopamine, in the periventricular and arcuate regions, and with CRH in the paraventricular nucleus. Some investigators have shown that it alters PRL release and blunts the response of that hormone to TRH [153]; it may also play a role in the regulation of GH and LH. Substance P

FIGURE 1.9. By electron microscopy, the normal human posterior lobe of pituitary contains numerous axonal processes filled with neurosecretory material (arrows). A large, dilated nerve terminal is the ultrastructural equivalent of the Herring body. (Magnification ¥7960)

evidence suggests that the brain is a site of glucagon biosynthesis. Neurons staining for glucagon and glucagon-like peptides have been detected in different regions of the brain including retina, hypothalamus, and medulla oblongata [145,146] and glucagon gene expression has been demonstrated in the brain stem and hypothalamus [147]. Intracerebral injections of glucagon produce dosedependent hyperglycemia, suggesting that this substance plays a role in the regulation of glucose metabolism centrally as well as peripherally [148]. Neuropeptide Y (NPY)

This 36-amino acid peptide is found by immunocytochemistry in a widely distributed pattern throughout the brain [149,150]. NPY-containing neurons are particularly numerous in the arcuate nucleus of the hypothalamus, striatum, hippocampus, and cortex. Nerve fibers are abundant in the paraventricular and suprachiasmastic nuclei and fibers are found in the internal zone of the median eminence. High

Substance P has been found in neurons in various regions of the brain. There are multiple substance P-containing afferents to the hypothalamus from the amygdala via the stria terminalis. Hypothalamic neurons containing this peptide are abundant in the arcuate, dorsomedial, ventromedial, and premamillary nuclei [152,154]. It is colocalized with vasopressin, is probably involved in pain perception within the limbic system, and may be secreted into the hypophysial portal vasculature where it is thought to affect blood flow. Substance P has been shown to alter GH regulation in vivo, but this is postulated to act via regulation of somatostatin neurons. It has variable effects on release of PRL, TSH, and LH. Bombesin/Gastrin-releasing Peptide (GRP)/Neuromedin

Bombesin was originally isolated from the skin of amphibians and stimulated great interest when it was shown to have potent biologic activity in mammals. Mammalian counterparts were soon discovered and given the names gastrin-releasing peptide and neuronedin. Bombesin-like immunoreactivity was found in neurons of the external zone of the median eminence near small blood vessels and colocalized with CRH [155]. Pharmacologic studies showed that bombesin can alter the release of adenohypophysial hormones including GH, PRL, and LH. The discovery of immunoreactivity for homologous peptides in hypothalamus suggests a possible role for these substances in the regulation of pituitary function. Calcitonin Gene-related Peptide (CGRP)

This 37-amino acid peptide is derived from the calcitonin gene by alternative messenger RNA (mRNA) processing.

Chapter 1

It is detected primarily in nervous tissue; CGRPimmunoreactive cell bodies are found in the supraoptic, paraventricular, and infundibular nuclei of the human hypothalamus [156]. CGRP inhibits GH release both in vitro and in vivo. It may also affect PRL release [157]. POMC-derived Peptides b-Endorphin

and other POMC-derived peptides are widespread in the central nervous system, their highest concentration being in the arcuate nucleus of the hypothalamus [158]. Nerve fibers containing b-endorphin and melanocyte-stimulating hormone-a (MSH-a) extend throughout many regions and innervate the median eminence, amygdala, preoptic area, ventromedial nucleus, and capillaries of the median eminence and portal system. There is another distinct and complex network of fibers containing the opiate methionine enkephalin (met-enkephalin). In addition to the multiple effects of these peptides on behavior, pain perception, immunomodulation, and many other functions too numerous to review here, they are also thought to modulate pituitary function [159], by inhibiting ACTH and LH release and stimulating PRL secretion. The effect on PRL is likely both direct and by reducing dopaminergic inhibition; the inhibition of LH appears to be only indirect via GnRH inhibition. The pituitary actions of endogenous opioids are complex, since they are themselves regulated by gonadal steroids and opioids may play a role in the negative-feedback effects of gonadal steroids on gonadotropins. Opioids also suppress the release of vasopressin and oxytocin, directly and possibly via noradrenergic regulation [160]. Cytokines

The close link between the neuroendocrine and immune systems has become widely recognized. Hormones are known to modulate the reactivity of immunocompetent cells and cytokines secreted by members of the immune system simulate secretion of hormones, most notably in the hypothalamus–pituitary–adrenal axis [161]. Interleukins are a family of peptides that mediate the immune response. Interleukin-1 (IL-1) is a monokine produced by macrophages in response to antigenic challenge. IL-1 stimulates ACTH and suppresses GH, LH and TSH secretion [162]. IL-1b immunoreactivity has been reported in neuronal fibers of the human brain, including the subfornical organ, stria terminalis, ventromedial nucleus of the hypothalamus, the posterior hypothalamus, and around the vessels of the median eminence. It is found in areas containing CRH-immunoreactive neurons and is known to increase CPH-mediated ACTH but is not found in the same synaptic vesicles as CRH. In fact, it is not released by depolarizing stimuli and it is not clear that IL-1b is even produced by neurons [161]. It also stimulates somatostatin biosynthesis in cultures of fetal rat brain [163], providing a

15

Functional Anatomy of the Hypothalamic Pituitary Axis

likely explanation for the in vivo reduction of GH and TSH secretion. Other interleukins are not known to be hypothalamic peptides but affect pituitary function. Interleukin-6 (IL-6) stimulates release of GH, PRL, and LH; tumor necrosis factor-a (TNF-a) and interferon-g also affect release of PRL and LH [164–166]. This information has significant neuroendocrinologic implications in the study of reproduction, autoimmunity, and other diseases. Growth Factors

A number of peptide growth factors, classified into several major families, are known to modulate cell proliferation. [167] Some are also known to affect hormone production (Table 1.1) and some are, in turn, modulated by hormones. Only a few have been identified in the hypothalamus and are thought to play a physiologic role in pituitary regulation; these include fibroblast growth factor (FGF) [168] and transforming growth factor-a (TGF-a) [169]. The former increases release of PRL and reduces basal GH; it also modifies the response of pituitary hormone release to TRH [170]. The latter has been localized by immunocytochemistry mainly in magnocellular neurons of the paraventricular and supraoptic nuclei of the hypothalamus. It is structurally homologous to epidermal growth factor (EGF) and may mediate some EGF effects on adenohypophysial production of GH, PRL, and TSH as well as cell proliferation [171].

Classification of Hypothalamic and Neurohypophysial Pathology Numerous pathologic processes involve the hypothalamus and neurohypophysis [46,47], giving rise to various symptoms, signs, and clinical syndromes. Among these, endocrine changes may be conspicuous. The local symptoms, such as visual defects, nerve palsy, and headache, may be secondary

Table 1.1. factors

Hormone regulation by pituitary-derived growth

Peptide

GH

PRL

EGF Activin FGF IGF-I IGF-II Insulin Endothelin Interleukin-1

Ø Ø

≠

Ø

FSH/LH

ACTH ≠

≠ ≠

Ø Ø Ø

TSH

≠

≠ ≠

≠ ≠

≠

GH, growth hormone; PRL, prolactin; TSH, thyroid-stimulating hormone; FSH, follicle-stimulating hormone; LH, luteinizing hormone; ACTH, adrenocorticotropic hormone; EGF, epidermal growth factor; FGF, fibroblast growth factor; IGF, insulin-like growth factor.

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to increased intracranial pressure or may be due to compression, invasion, injury, or destruction of tissue components. Involvement of certain parts of the hypothalamus can give rise to nonendocrine pathology, including abnormalities of appetite and temperature regulation. The endocrine sequela of hypothalamic disorders can be classified into two categories: (i) those giving rise to hypothalamic hormone deficiency; and (ii) those giving rise to hypothalamic hormone excess. The major pathologic processes causing these disorders are reviewed briefly and their features are fully discussed in ensuing chapters. Hypothalamic Hormone Deficiency

Insufficient secretion of hypothalamic adenohypophysiotropic hormones may be caused by destructive lesions of the hypothalamus. Lesions such as granulomas, meningitis or other inflammatory processes, pituitary adenomas, craniopharyngiomas, lymphomas, histiocytosis-X, metastatic carcinomas, craniocerebral trauma or hypoxic tissue injury may impair the synthesis of numerous hormones, including vasopressin, oxytocin, and the various adenohypophysiotropic hormones. These lesions give rise to true hypothalamic hypopituitarism. Similarly, lesions affecting the pituitary stalk, such as surgical or traumatic sectioning of the hypophysial stalk or its interruption by various pathologic processes, block the transport of the hypothalamic hypophysiotropic hormones from their sites of production in the hypothalamus to the anterior pituitary. It is obvious that any disconnection of the direct link between the hypothalamus and pituitary results in various degrees of hypopituitarism. Hypocortisolism, hypogonadism, and/or hypothyroidism have been demonstrated conclusively in patients with destructive diseases of the hypothalamus. Deficiency of GHRH usually results in growth retardation or in dwarfism [172]. Destructive lesions may manifest as growth retardation of hypothalamic origin in which patients respond to administration of GHRH. Idiopathic GH deficiency may be associated with other pituitary insufficiency and hypothalamic hormone abnormalities. Of particular interest is an autosomal recessive form of hypothalamic dwarfism due to failure to produce GHRH [173,174]; this disorder represents approximately 10% of cases of inherited growth retardation. The limited morphologic studies of patients with GHRH deficiency have documented that pituitary somatotrophs have normal morphology and GH content and administration of GHRH elicits a normal GH response [175]. Destructive lesions of the hypothalamus may impair the synthesis and/or release of dopamine and therefore may give rise to hyperprolactinemia [96]. Elevated blood PRL levels are also found in patients with disruption of, or damage to, the pituitary stalk. As a result of decreased transport of dopamine, lactotrophs are relieved from their tonic hypothalamic inhibition, leading to hyperprolactinemia. Hypothalamic hypothyroidism is usually not an isolated deficiency; it is most often associated with multiple hypo-

thalamic hormone deficiencies caused by organic hypothalamic pathology. Hypothalamic hypogonadism may be attributed to lack of GnRH or to nonpulsatile release of that hormone [176]. It may be an isolated defect of GnRH neurons [177]; in patients with hemochromatosis it is attributable to a combination of iron-induced damage to GnRH neurons and pituitary abnormalities [178]. Kallmann’s syndrome is an uncommon variant of hypothalamic hypogonadism which occurs in both men and women. The hypogonadism is associated with anosmia [179]. These deficits are due to defective development of the GnRH-secreting part of the hypothalamus associated with hypoplasia or aplasia of the olfactory area of the brain that can be documented by magnetic resonance imaging (MRI) [180]. Kallman’s syndrome is now known to be attributed to deletion or mutations of the KAL gene, located at Xp22.3; this gene encodes a 680 amino acid protein, anosmin, that has homology to other molecules implicated in neural development [181]. The pathology of the hypothalamus in this disorder has been thoroughly described in a histologic study in which abnormalities were found in various hypothalamic nuclei [182]; the lateral tuberal nuclei were underdeveloped and the subventricular nucleus exhibited neuronal hypertrophy. The number of pituitary gonadotrophs was markedly decreased as documented by immunohistochemistry and electron microscopy [182]. Detailed pathologic examination of the hypothalamus is yet to be reported in most patients with no major organic hypothalamic disease and apparently idiopathic deficiency of hypothalamic adenohypophysiotropic hormones. Diabetes insipidus is an uncommon but distinct clinical syndrome characterized by polyuria and polydipsia and results from vasopressin deficiency [46,47]. The extent of disease varies with the location of tissue destruction. Because vasopressin is transported through the pituitary stalk to the posterior lobe where it is stored in nerve endings, destruction of the posterior pituitary may interfere with storage and release of the hormone, resulting in its deficiency. This situation is usually transient and selective lesions of the posterior lobe usually cause only mild and temporary polyuria and polydipsia. Destruction high in the infundibulum results in permanent diabetes insipidus due to retrograde axonal degeneration and atrophy of neurons in the supraoptic and paraventricular nuclei [183]. Destruction of the supraoptic and/or paraventricular nuclei gives rise to permanent disease; approximately 90% destruction is required to cause significant and permanent disease [184], explaining why section low in the stalk, which creates atrophy of a minority of neuronal axons, causes only a transient disorder. Rare forms of hereditary idiopathic diabetes insipidus are usually unassociated with gross destruction of the hypothalamus [185]; detailed histologic studies reveal severe atrophy of neurons in the supraoptic and paraventricular nuclei in most patients but the etiology of these disorders remains unknown. In a few instances, neurons of the

Chapter 1

two magnocellular nuclei have been microscopically normal but devoid of vasopressin [186]. Some studies have suggested that this disorder may in some cases represent autoimmune destruction of vasopressin-producing neurons [187]. Oxytoxin deficiency, resulting from a decrease in the production of oxytocin, causes no major clinical abnormalities. Hypothalamic Hormone Excess

Hypersecretion of hypothalamic hormones, although rare, can result in endocrine symptomatology. Certain endocrine diseases characterized by hyperfunction of the anterior pituitary may be due primarily to adenohypophysiotropic hormone excess as a result of excessive secretion of hypothalamic releasing hormones. As yet, there is no evidence that hypersecretion of hypothalamic inhibiting hormones causes pituitary hypofunction. Acromegaly, a syndrome resulting from sustained GH hypersecretion, is almost always the result of a pituitary adenoma. Only one case of proven ectopic GH production has been reported [188]. More numerous are cases of production of GHRH by tumors that result in somatotroph hyperplasia or adenoma in the pituitary and produce clinical features of GH excess [87]. Extrahypothalamic GHRH production has been documented in endocrine tumors of lung, pancreas, thymus, medullary thyroid carcinoma, and pheochromocytoma. Primary tumors of the hypothalamus, called variously hypothalamic neuronal hamartomas, hypothalamic gangliocytomas and adenohypophysial neuronal choristomas, are lesions consisting of neurons with hypothalamic differentiation. Some of these tumors have been shown to contain GHRH and have been associated with pituitary somatotroph adenoma and acromegaly [189]. Several have been shown to be plurihormonal, containing GnRH, glucagon, gastrin, and somatostatin. Gigantism with pituitary GH excess has been described occasionally in the McCune–Albright syndrome [190]; this disorder is characterized by polyostotic fibrous dysplasia, pigmentary abnormalities and a variety of endocrine abnormalities, most commonly precocious puberty in female children. The pituitary may show mammosomatotroph hyperplasia that is attributed to activation of the signal transduction pathway generating cyclic AMP in these cells. These various manifestations are now known to be due to activating mutations of the Gs alpha gene (gsp mutations) [191,192]. Mammosomatotroph hyperplasia has also been reported as a cause of gigantism in the absence of McCune–Albright syndrome [193]. No proof of GHRH excess, either of hypothalamic or ectopic origin, has been offered in these patients; however, the pathogenetic role of GHRH in mammosomatotroph proliferation has been proven in mice transgenic for GHRH who have mammosomatotroph hyperplasia resembling that seen in these human disorders [194]. Cerebral gigantism (Sotos’ syndrome) has also been postulated to be a primary hypothalamic endocrinopathy. However, in patients with this

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disorder, hormone levels are normal and a morphologic study of the hypothalamus and pituitary showed no specific abnormality [47]. Cushing’s disease is known to be associated with pituitary corticotroph hyperplasia in some cases [195]; it has been suggested that the corticotroph hyperplasia may be secondary to excess production of CRH [196]. Ectopic production of CRH has been shown with increasing frequency and can cause Cushing’s disease; it must be considered in the differential diagnosis of that disorder [197,198]. In some cases, CRH has been elaborated in association with ACTH, suggesting autocrine regulation of some cases of ectopic ACTH syndrome. Rarely, a hypothalamic CRH-producing neuronal tumor has been reported in association with pituitary corticotroph hyperplasia or adenoma and Cushing’s disease [196,199–201]. Precocious puberty is an uncommon clinical syndrome which is in most instances idiopathic or attributable to androgen- or estrogen-producing neoplasms. Idiopathic precocious puberty is assumed to be hypothalamic in origin despite the absence of any anatomic lesion; less than 10% of cases are due to demonstrable hypothalamic pathology. Electroencephalographic abnormalities and behavioral disturbances have been observed in otherwise neurologically normal patients with this disorder [202]. Some patients respond to therapy directed against hypothalamic stimulation, but the disease may also be GnRH independent [203]. Structural lesions in and around the hypothalamus associated with precocious puberty fall into two groups. Some lesions actively participate in the production of precocious puberty by elaborating GnRH; these consist primarily of hypothalamic neuronal hamartomas which have been shown to contain GnRH in some cases [204,205]. The second group consists of hormonally inactive lesions which nonspecifically stimulate or inhibit hypothalamic centers involved in the regulation of sexual maturation. These are primarily lesions in the region of the posterior hypothalamus and include tumors of the pineal and hypothalamus, astrocytic gliomas, infections, hydrocephalus, head trauma, arachnoid cysts, and, occasionally, craniopharyngiomas. Precocious puberty is a prominent feature of the McCune–Albright syndrome [200]; in this disease, there is no demonstrable hypothalamic lesion and the patients may develop normal sexual function in adulthood. The hypothalamus has been implicated as the cause of some cases of hyperthyroidism but this has not been proven [206]. Vasopressin excess may cause water intoxication. The syndrome of inappropriate antidiuretic hormone (ADH) secretion (SIADH) may be due to idiopathic hypothalamic vasopressin excess or may be associated with various disease, including malignant tumors, most often bronchogenic carcinoma, and tuberculosis [207]. In the latter instances, this syndrome may be due to secretion of vasopressin or a similar peptide by tumoral or diseased pulmonary tissue [208,209]. In patients in whom no underlying cause is found, the

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idiopathic syndrome is likely due to posterior pituitary hyperfunction but no morphologic characteristics of this disorder have been described. The pathologic process in the brain may be in a number of areas; inhibitory osmoreceptors are found diffusely in the brain, as far away as the brain stem, and lesions can reduce the tonic inhibition of vasopressin, thereby increasing its release. Evidence of this widespread control is found in the increase in vasopressin release associated with congestive heart failure, cirrhosis, renal disease, brain trauma, positive pressure ventilation, and neurosurgical manipulation of the pituitary and hypothalamus. Age Related Changes in the Hypothalamus

Age related changes in the hypothalamus and neurohypophysis are few. The most striking is a morphologic curiosity, the accumulation of corpora amylacea, most often seen in the region of the supraoptic nuclei. The magnocellular nuclei are remarkably unaffected by age; immunohistochemistry shows no significant change in staining of vasopressin and oxytocin in older subjects [210]. The volume of the mamillary bodies is reduced but cell numbers appear normal [211]. Lipofuscin accumulation is commonly seen in the aging brain but does not appear to affect hypothalamic neuronal function. The suprachiasmatic nucleus decreases in volume and cell number with age [53]. The subventricular nucleus, in contrast, undergoes marked hypertrophy of its usually parvicellular neurons in postmenopausal women, as well as in starvation, posthypophysectomy, late pregnancy, and in patients suffering from prolonged postpartum hypopituitarism [58–64]. ADENOHYPOPHYSIS

Anatomy The adenohypophysis is composed of three parts, the pars distalis, the pars intermedia, and the pars tuberalis (Fig.

(a)

1.10). The pars distalis constitutes the largest portion of the gland; it is also known as the anterior lobe or the pars glandularis. The pars intermedia is rudimentary in the human pituitary; it is the vestigial posterior limb of Rathke’s pouch and is found in its underdeveloped form behind the medial cleft of the pituitary. It is also known as the intermediate lobe. The pars tuberalis is an upward extension of a few layers of adenohypophysial cells surrounding the external aspect of the lower hypophysial stalk; it is also known as the pars infundibularis. The adult human pituitary is bean shaped, bilaterally symmetrical, and measures approximately 13 mm transversely, 9 mm anteroposteriorly, and 6 mm vertically. It weighs approximately 0.6 g. The anterior lobe comprises about 80% of the pituitary. The pituitary of women is somewhat larger than that of men; moreover, the pituitary is heavier in multiparous than in nulliparous women. The increased size is attributable to the marked hyperplasia of the pituitary during pregnancy and lactation, which may increase the weight to 1 g or more; postlactational involution occurs but the gland does not return to its prepregnancy size. A slight to moderate weight reduction is seen with advancing age. The hypophysis is found within the sella turcica where it is lined by dura mater, a dense layer of connective tissue. The uppermost portion of the adenohypophysis is covered by the diaphragma sellae, a reflection of the dura that constitutes the roof of the sella turcica. The sellar diaphragm has a small central opening for the hypophysial stalk and protects the pituitary from the pressure of cerebrospinal fluid (CSF). Defective development or absence of this structure may lead to the so-called empty sella syndrome in which the pituitary is compressed by CSF pressure. The increased pressure results in enlargement of the sella turcica and flattening of the pituitary, which may be compressed to a thin layer of tissue at the bottom of the sella turcica. This lesion is usually unassociated with functional

(b)

FIGURE 1.10. (a) Sagittal section of an adult human pituitary obtained at autopsy illustrates the adenohypophysis (A) and neurohypophysis (N) attached to the pituitary stalk. (b) Horizontal cross-section of an adult human autopsy pituitary allows accurate identification of the neural lobe (N) and the anterior lobe (A) with subtle distinction between the two lateral lobes and the median wedge. The cystic remnants of Rathke’s cleft that represent the pars intermedia in humans are indistinct (arrow head).

Chapter 1

hypophysial abnormalities [206] but is important in differential diagnosis. Other anatomic variations in the shape of the hypophysis, the size and width of the sellar diaphragm, and relation to pituitary structures have been reported [207]. Minor changes have no endocrine significance.

Vascular Supply The blood circulation of the human hypophysis has major implications for the regulation of adenohypophysial hormone secretion [48–51]. The gland receives blood through the hypophysial portal circulation which carries the hypothalamic stimulatory and inhibitory hormones from the infundibulum to adenohypophysial cells. As described above (see Fig. 1.2), the branches of the superior hypophysial arteries penetrate the stalk, form a superficial network of vessels (external plexus) and give rise to a mesh of capillaries and gomitoli (internal plexus). The latter presumably regulate blood flow, thereby affecting transport of hypothalamic hormones to the adenohypophysis. The long portal vessels that arise from the infundibular plexuses and the short portal vessels that originate in the lower pituitary stalk and posterior lobe enter the pars distalis carrying adenohypophysiotropic hormones; the former carry 70–90% of the blood flow while only 10–30% originates in the short portal vessels. The trabecular or loral arteries transport blood to the adenohypophysis via the neural lobe, but, in addition, some arterial blood is directed to the adenohypophysis via two branches of the inferior hypophysial artery: (i) the capsular artery, which serves the connective tissue of the pituitary capsule and penetrates to the superficial cell rows of the adenohypophysis; and (ii) the artery of the fibrous core. In some individuals, the middle hypophysial artery may vascularize the adenohypophysis directly [212]. The volume of veins leading away from the adenohypophysis is considerably smaller than that of portal vessels entering the gland. The meager venous connections drain adenohypophysial and neurohypophysial blood to the cavernous sinus. It has been shown that the short portal vessels also serve as efferent channels, giving rise to reversal of blood flow in the neurohypophysial vascular bed. This implies that secretory products of the adenohypophysis may also enter the neurohypophysis and, by reverse flow to the median eminence, play a role in the regulation of hypothalamic factors via blood flow [50,51]. Pituitary capillaries are lined by fenestrated endothelium with a thin subendothelial space. Hormones released by adenohypophysial cells must therefore pass through the basement membrane of their cell of origin, capillary basement membrane, subendothelial space, and the endothelial cell layer to reach the bloodstream.

Nerve Supply The adenohypophysis has no direct nerve supply, apart from small sympathetic nerve fibers which are associated with and

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presumably innervate capillaries [195]. In some species, the intermediate lobe has rich innervation [213]. Thus, although neural connections may affect blood flow to the adenohypophysis, it is unlikely that they are involved in the regulation of adenohypophysial hormone secretion, which is predominantly neurohumoral.

Embryology All three portions of the adenohypophysis are assumed to derive from Rathke’s pouch, an endodermal pouch of the primitive oral cavity. At the third week of gestation, endoderm from the roof of the stomodeum thickens and begins to invaginate; by 5 weeks, Rathke’s pouch is a long tube with a narrow lumen and a thick wall composed of stratified cuboidal epithelium. By 6 weeks, the connection with the oropharynx is totally obliterated and Rathke’s pouch establishes direct contact with the downward extension of the hypothalamus that gives rise to the infundibulum. The two tissues are enclosed by the cartilage anlage of the sphenoid bone, separating them from the stomodeum; the sella turcica is formed by 7 weeks [66]. It was suggested that Rathke’s pouch arises from the ventral neural ridge in the pharyngeal region, thus sharing with the hypothalamus and posterior pituitary a common neuroectodermal origin [214–217]. Some features of adenohypophysial cells suggest that they are members of the APUD (amine precursor uptake and decarboxylation) cell system. The use of avian allografts, biologic markers, and serial sections of early chick embryos has provided indirect evidence for this theory; however, further proof is required for its validation. As the cells of Rathke’s pouch proliferate, the anterior portion forms the pars distalis and pars tuberalis whereas the posterior wall lies in direct contact with the posterior lobe anlage and becomes the pars intermedia [218,219]. The growth of the anterior limb extends laterally and follows a triradiate pattern; the lateral borders become the lateral wings of the adult gland and the midline portion becomes the anteromedial “mucoid wedge.” By midgestation, the medial cleft becomes a residual lumen and growth of the pars nervosa reverses the convexity of the posterior wall of the cleft to a concave structure. The border between Rathke’s pouch and the pars nervosa becomes indistinct; it consists of remnants of the obliterating lumen, and a few cystic cavities lined by cuboidal or columnar epithelium. This represents the rudimentary pars intermedia of the human hypophysis. The pituitary grows rapidly in early fetal life: the mean weight at 10–14 weeks of gestation is 3 mg, at 25–29 weeks 50 mg, and at term approximately 100 mg [218]. The pituitary portal vascular system begins to form before 7 weeks of gestation, and by 12 weeks the anterior pituitary and median eminence are well vascularized. Portal vessels are recognized at 11.5–14 weeks, are well developed by 15–16 weeks, and are fully established by 18–20 weeks [71,72].

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The first cell type to develop in the human fetal pituitary is the corticotroph [219–221]; at 6 weeks of gestation, cells with ultrastructural features of differentiating corticotrophs are found, and by 7 weeks, ACTH immunoreactivity is detectable. At 8 weeks of gestation, somatotrophs are identified by ultrastructural criteria and contain intense GH immunoreactivity. a-Subunit of the glycoprotein hormones is found in cells which have features of the glycoprotein hormone cell line; differentiated thyrotrophs and gonadotrophs are found subsequently at 12 weeks when b-subunits of those hormones are immunolocalized. Gonadotrophs show sexual dimorphism in the fetus; the differences are most marked from 15–25 weeks when this cell type is more numerous in female glands. Throughout gestation, LH-containing cells predominate in male glands, unlike the more equal distribution of LH- and FSHcontaining cells in female tissues [220]. Lactotrophs are the last cells to differentiate in the human fetal pituitary. PRL is found only at 12 weeks of gestation and is localized in mammosomatotrophs, bihormonal cells which appear to be the sole source of PRL until 24 weeks of gestation [221]. Differentiated lactotrophs are found after that time and undergo a striking hyperplasia in the late third trimester, analogous to that seen in late gestation and lactation in the pituitary of the mother. Novel transcription factors that play a role in anterior primordial development are being identified at a rapid pace. Many of these are implicated in early pituitary organogenesis, including the bicoid-related pituitary homeobox factor Ptx1 [222], Pituitary homeobox factor 2 (Ptx2), structurally related to Ptx1 [223], two members of the Lhx gene family, a group of LIM homeobox genes, Lhx3 and Lhx4 [224] and P-LIM, another LIM homeobox protein transcription factor that is selectively expressed in the pituitary with highest levels at the early stages of Rathke’s pouch development [225]. Another early marker of pituitary differentiation is the Rathke’s pouch homeobox (Rpx) protein which is identified in the pituitary primordium prior to the onset of known pituitary hormone production [226]. The Prophet of Pit-1 (PROP-1) is a paired-like homeodomain protein that is expressed early in pituitary development. It induces Pit-1 expression and plays a role in downregulation of Rpx [227,228]. Inactivating mutations of PROP-1 have been identified as the cause of Pit-1 deficiency in Ames dwarf mice [227] and in humans with combined pituitary hormone deficiency [229,230]. Id, a member of the helix-loop-helix (HLH) family of transcription factors is also found early in development and in some pituitary tumor cell lines but is decreased or absent in differentiated cells [231]. Its role in pituitary cytodifferentiation remains unclear. The molecular factors that determine hormone production have now been identified as transcription factors that target specific hormone genes. These factors have clarified three main pathways of cell differentiation [232,233]. ACTHproducing corticotrophs are determined by a novel T box

factor in cooperation with Ptx1 and corticotropin upstream transcription-binding element (CUTE) proteins including neuroD1/beta2 [234,235]. Bihormonal gonadotrophs require expression of steroidogenic factor (SF)-1 [236,237]. Pit-1 [238] directs differentiation of a complex family of cells that can mature into somatotrophs, mammosomatotrophs, lactotrophs or thyrotrophs [239–241] with the additional expression of estrogen receptor (ER) a [242], which enhances PRL secretion, or thyrotroph embryonic factor (TEF) which stimulates TSH-beta production [243]. The recognition of these molecular determinants of adenohypophysial cytodifferentiation has clarified the patterns of plurihormonality which have been recognized in pituitary adenomas and provide a framework for classification of these tumors. In rodents and humans, differentiation and/or maintenance of somatotroph, lactotroph and thyrotroph phenotypes are dependent on expression of a functional pit-1 gene; mutations in the pit-1 gene result in hypopituitarism [244–247] and hypoplasia of somatotrophs, lactotrophs and thyrotrophs [244]. An interesting observation is that Pit-1 mRNA and protein are highly expressed during human pituitary development at 17–19 weeks, when GH levels are extremely high, and near term when there is proliferation of lactotrophs [248]. These data suggest that Pit-1 plays an important role not only in the differentiation process, but also in the regulation of hormonal activity and possibly also of cell proliferation. Gonadotroph differentiation likewise requires SF-1; mice with disruption of SF-1 fail to develop pituitary gonadotrophs [236] as well as a ventromedial nucleus of the hypothalamus and steroidogenic glands [249].

Functional Anatomy In contrast with the hypothalamus, the cell types of the adenohypophysis are highly characterized with respect to structure and function. It is notable that with the exception of ACTH and a-subunit of the glycoprotein hormones, ectopic production of pituitary hormones is rare. Somatotrophs

Approximately 50% of the cells of the adenohypophysis produce GH. Most of these are located in the lateral wings of the anterior lobe; scattered somatotrophs are found in the median wedge [195]. These cells can usually be identified by light microscopy as medium-sized acidophilic cells that stain with eosin, phloxin, and orange G. They are spherical or oval in shape, with central, spherical nuclei. Immunohistochemistry reveals intense positivity for GH distributed throughout the cytoplasm [38] and for Pit-1 in the nucleus [221,250]. Occasionally, smaller cells contain GH positivity in a globular structure which represents the Golgi complex; these may be sparsely granulated, actively secreting cells. In situ hybridization localizes GH mRNA to both densely granulated acidophils and to occasional chromophobes (Fig. 1.11). By electron microscopy, somatotrophs are spher-

Chapter 1

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FIGURE 1.11. In situ hybridization localizes growth hormone messenger RNA in the normal pituitary in large cells with acidophilic cytoplasm and in occasional chromophobes. (Magnification ¥102)

ical or oval cells with centrally located spherical nuclei and cytoplasm of relatively low electron density [40]. The prominence of rough endoplasmic reticulum and Golgi regions varies with the secretory activity of the cell; active cells generally have well-developed, lamellar rough endoplasmic reticulum, and a large Golgi complex, whereas less active ones tend to have less conspicuous synthetic organelles. In the majority of somatotrophs, the cytoplasm is occupied by spherical, evenly electron-dense secretory granules which store hormone (Fig. 1.12). The limiting membrane of the granules is tightly opposed. The secretory granules range widely in size from 150 to 800 nm in diameter with an average of 350–500 nm. Secretory granules may be found accumulated at the cell periphery but granule extrusions are not detected. Ultrastructural immunocytology confirms the presence of GH in secretory granules and within the Golgi region of actively secreting cells. A subset of these cells has been recognized to contain a-subunit of glycoprotein hormones [40]. The incidence, distribution, morphology, and hormone content of somatotrophs are remarkably constant in the human pituitary. They do not appear to be affected by age, sex, various disease states, or drug therapies. Even in glands containing GH-producing adenomas the nontumorous somatotrophs show no evidence of suppression and cannot be distinguished from those in normal glands [40]. An exception to this morphologic uniformity is the case of long-standing hypothyroidism in which some degranulation of somatotrophs may occur; this change is much less marked than the almost complete degranulation of somatotrophs due to hypothyroidism which has been documented in rodents [250]. In pituitary adenomas, somatotrophs display a striking dimorphism [195]. Despite the morphologic differences between densely granulated and sparsely granulated soma-

FIGURE 1.12. A nontumorous somatotroph contains short profiles of rough endoplasmic reticulum (arrow head), a moderately developed Golgi complex (G) and numerous spherical, evenly electron-dense secretory granules with tightly fitted limiting membranes. (Magnification ¥5500)

totroph adenomas, both types have similar clinical features with elevated blood GH levels and acromegaly. Although the GH concentration in the tumor is proportional to the degree of granularity, neither of these correlate with blood GH levels. The morphine variant does, however, predict response to octreotide therapy [251]. Densely granulated somatotroph adenomas are composed of intensely acidophilic cells which contain strong immunoreactivity for GH throughout their cytoplasm. The tumor cells resemble resting nontumorous somatotrophs, with welldeveloped rough endoplasmic reticulum, prominent Golgi complexes and numerous spherical, evenly electron-dense secretory granules which measure 300–600 nm in diameter. These tumors may contain immunoreactivity for prolactin and/or a-subunit of glycoprotein hormones. The presence of these substances and GH in the same secretory granules has been documented by ultrastructural immunocytology using the double immunogold technique [252]. The use of keratin immunocytochemistry distinguishes these cells from

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FIGURE 1.13. A sparsely granulated adenomatous somatotroph harbors a characteristic juxtanuclear fibrous body composed of intermediate filaments that trap smooth endoplasmic reticulum, Golgi saccules (arrowheads), centrioles (arrows) and pleomorphic forming secretory granules. (Magnification ¥11390)

other somatotroph adenomas. Densely granulated cells have perinuclear low molecular weight cytokeratin filaments. Sparsely granulated somatotroph adenomas are composed predominantly of chromophobic or only slightly acidophilic cells which may show considerable nuclear pleomorphism. The nuclei are generally concave and binucleate; multinucleate cells are easily found. By light microscopy, spherical light areas can be seen adjacent to the nuclei; these correspond to the fibrous bodies seen by electron microscopy and are known to contain immunoreactive keratin [254]. The application of cytokeratin staining allows recognition of fibrous bodies that characterize this tumor [253]. GH positivity is found generally in the juxtanuclear Golgi region, while secretory granules, which are few, can be seen usually only with oil immersion. By electron microscopy, this tumor has a highly characteristic ultrastructure which differs significantly from that of nontumorous somatotrophs (Fig. 1.13). The tumor cells are irregularly shaped with numerous indentations and have lobulated pleomorphic and eccentrically located nuclei. The globular cytoplasmic

fibrous body, found in up to 95% of cells in sparsely granulated somatotroph adenomas, consists of intermediate-sized filaments that accumulate in the Golgi region, close to the concave side of an indented eccentric nucleus. The nearly perfectly globular structure traps tubular smooth endoplasmic reticulum, Golgi saccules, centrioles, mitochondria, lysosomes, and secretory granules. The secretory granules in the cells are scattered throughout the cytoplasm and are smaller than in densely granulated cells, measuring 100 – 250 nm in diameter. No exocytoses are seen. GHRH has been shown to stimulate proliferation of GH-containing cells in vitro during long-term incubations [255]. The effects of chronic exposure to GHRH in humans can be deduced from the study of pituitaries of patients with tumors producing GHRH [40,87]. These patients have marked somatotroph hyperplasia; the somatotrophs are generally large with very well-developed rough endoplasmic reticulum profiles and large Golgi complexes. The hyperplastic cells have numerous large electron-dense secretory granules characteristic of nontumorous cells; sparsely granulated cells with fibrous bodies are generally not seen. Although short-term exposure to GHRH is known to deplete the GH pool [256] and to convert densely granulated cells to more sparsely granulated forms [257], cells of densely granulated somatotroph adenomas chronically exposed to GHRH in vitro show an increase in the cytoplasmic volume densities of rough endoplasmic reticulum and Golgi complexes and a decrease in cytoplasmic volume density of secretory granules. However, they do not assume the characteristics of sparsely granulated cells and do not contain fibrous bodies [258]. Sparsely granulated adenomas undergo similar alterations during long-term exposure to GHRH in vitro and retain their characteristic fibrous bodies [258]. Although GHRH stimulates somatotroph proliferation in patients with GHRH-producing tumors, the hyperplastic adenohypophysial cell population in mice transgenic for this hormone is dominated by mammosomatotrophs as well as somatotrophs [194]. The presence of mammosomatotrophs may be attributed to exposure to GHRH excess in these animals during fetal life, at which time mammosomatotrophs are numerous, in contrast to the usual onset of tumoral GHRH excess in adulthood. Further studies are required to clarify the complex regulation of somatotroph and mammosomatotroph differentiation and proliferation by GHRH. The morphologic effects of somatostatin exposure on nontumorous somatotrophs has not been documented. The use of somatostatin analogs, such as SMS 201–995 (Sandostatin or Octreotide) in the therapy of acromegaly has allowed morphologic study of the effects of that substance on somatotroph adenoma cells. Both in vivo [259–261] and in vitro [262], somatostatin and its long acting analogs induce inconsistent morphologic alterations. In some tumors, there is no change, while in others, cell size is reportedly reduced. In some cases, there is an increase in the cytoplasmic volume density of lysosomes unassociated with significant alteration

Chapter 1

of other subcellular organelles. A few studies have indicated that these substances may induce fibrosis; others have found vascular changes which may cause cell necrosis and tumor shrinkage but direct cytotoxic effects have not been documented in vitro. No inhibition of hormone synthesis has been identified, as indicated by reduction in the cytoplasmic volume density of rough endoplasmic reticulum or Golgi complexes; this suggestion confirms molecular studies which have shown no suppression of mRNA in somatostatinexposed nontumorous adenohypophysial cells [263] or tumors [264]. The morphologic studies suggest that somatostatin suppresses hormone release and may stimulate lysosomal degradation of stored hormones. There is, however, differential response to octreotide in vivo by the two somatotroph adenoma types; densely granulated tumors exhibit greater reduction of growth hormone secretion than do sparsely granulated tumors [251]. Lactotrophs

Lactotrophs, or PRL cells, represent approximately 15% of the cells of the adenohypophysis. However, the number of these cells shows wide variation, related to age, sex, and parity in women. In adult men and nulliparous women they constitute approximately 9% of adenohypophysial cells, whereas in multiparous women they represent up to 31% of the cell population [266]. PRL cells are randomly distributed throughout the anterior lobe but are most numerous in the posteromedial portions, with large numbers extending to the posterolateral aspect [195]. Using conventional stains, PRL cells are acidophils that are indistinguishable from somatotrophs; however, Herlant’s erythrosin and Brookes’ carmoisine allow selective visualization of densely granulated lactotrophs, although these techniques are inconsistent and are not

FIGURE 1.14. Immunocytochemistry identifies prolactin diffusely in the cytoplasm of polyhedral and elongated densely granulated cells (arrow), and in the juxtanuclear globular Golgi region of sparsely granulated cells (arrow head). (Magnification ¥102)

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sufficiently sensitive to detect sparsely granulated forms. Immunohistochemistry reveals larger numbers of PRLcontaining cells among which two populations can be distinguished [38] (Fig. 1.14). Densely granulated, polyhedral, or elongated cells are frequently found close to capillaries randomly distributed throughout the anterior lobe; they have abundant cytoplasm almost completely filled with dense granular positivity corresponding to secretory granules. Sparsely granulated cells, in contrast, are more numerous and are found predominantly in clusters at the posterolateral portion of the gland. They are elongated or angular cells with long cytoplasmic processes and strong immunoreactivity for PRL in a juxtanuclear globular Golgi complex. It has been postulated that the densely granulated cells store PRL, whereas the sparsely granulated cells are actively secreting forms. In addition to nuclear Pit-1 reactivity these cells may also exhibit staining for estrogen receptor a [265a,265b]. By electron microscopy, the two cell types are readily distinguishable [40] (Fig. 1.15). Densely granulated cells are

FIGURE 1.15. A nontumorous lactotroph has abundant cytoplasm with highly developed rough endoplasmic reticulum in parallel arrays (*) and few, small secretory granules. (Magnification ¥6080)

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rare in the adult pituitary but are more common in childhood and adolescence. They have ovoid or elongated cell bodies, well-developed rough endoplasmic reticulum at the cell periphery and a spherical or oval Golgi complex containing a few forming granules. The cell cytoplasm is almost completely filled with spherical, oval, or irregularly shaped large granules that have evenly electron-dense cores and measure up to 650 nm in diameter. Granule extrusions are seen occasionally, usually on the cell surfaces overlying basement membrane. The majority of lactotrophs in the adult gland are sparsely granulated, elongated or polygonal cells which may have multiple cell processes extending from the center of acini to the basement membrane and intimately surrounding gonadotrophs. They contain richly developed rough endoplasmic reticulum found in parallel arrays and occasionally forming concentric structures known as “Nebenkern” formations. The Golgi apparatus is prominent and contains pleomorphic immature secretory granules. Scattered secretory granules are fewer and smaller than in storing cells; the diameters range from 150 to 250 nm. Granule extrusions are common and are found not only at the basal cell surface but also on the lateral cell borders, distant from capillaries and basement membranes; the term “misplaced exocytosis” is used to designate this form of granule extrusion. Ultrastructural immunocytology localizes PRL in secretory granules of both cell types; cloudy positivity can also be found in the Golgi region of sparsely granulated forms. In the fetus [220] and newborn as well as during pregnancy and lactation [266,267] there is marked hyperplasia of PRL cells which has been attributed to stimulation by estrogens. During pregnancy and lactation the weight of the pituitary gland may increase to more than 1 g and almost 50% of the total pituitary cell population is composed of PRLcontaining cells. Exogenous estrogens are known to induce only mild hyperplasia of lactotrophs in the human pituitary; the increased cell population is composed predominantly of the small sparsely granulated cell type [268a]. With sustained stimulation of PRL synthesis due to decreased dopamine inhibition or increased estrogen levels, there is a progressive increase in cytoplasmic volume and accumulation of highly organized rough endoplasmic reticulum, enlarged Golgi, numerous forming granules, and few small stored granules [40]. In animals lacking the dopamine D2 receptor, there is extensive lactotroph hyperplasia that progresses to adenoma [268b]. Suppressed lactotrophs are found in some pituitary glands containing PRL-producing adenoma or in patients treated with dopamine agonists. These inactive cells have a reduced nuclear volume, irregular indented nucleus with coarse clumped heterochromatin, small cytoplasm with poorly developed organelles, and secretory granules which measure 50–300 nm; the granules have an increased relative cytoplasmic volume due to the markedly reduced total cytoplasmic volume. Occasional exocytoses may be found at the basal cell surface. These suppressed lactotrophs are identified by immunocytology at the electron microscopic level; they

cannot be characterized on the basis of their ultrastructural features alone, which are not unlike those of null cells. PRL cell adenomas also occur in densely and, more commonly, in sparsely granulated varieties [195]. There is no correlation between morphology and blood PRL levels, age, or sex of the patient. The cells resemble the densely and sparsely granulated cells of the nontumorous pituitary. Effects of dopamine agonists on the cells of lactotroph adenomas include significant reduction of cytoplasmic, nuclear, and nucleolar areas [269]. There is also a marked reduction in the cytoplasmic volume densities of rough endoplasmic reticulum and Golgi complexes, whereas secretory granules increase in size and cytoplasmic volume density. The size reduction of bromocriptine-treated prolactinomas is therefore attributable to reduction of individual cell size rather than to a decrease in cell number. Protracted therapy may result in fibrosis which can adversely affect surgical outcome [270]. Mammosomatotrophs

The existence of bihormonal cells containing both GH and PRL in the nontumorous human pituitary has been recognized only in the last few years [40]. This cell type was initially identified in pituitary tumors [195]; it was subsequently found to be the cell of origin of PRL in the fetal pituitary during the first half of gestation [221]. Mammosomatotrophs are thought to be randomly distributed throughout the anterior lobe. They cannot be recognized using conventional histologic techniques; they are acidophils and are indistinguishable from somatotrophs. Immunohistochemistry reveals intense GH content within these densely granulated, polyhedral cells. PRL content is also identified but staining is usually less intense. Nuclear Pit-1 and ER staining is usually strong [239,240,265a,265b] and these cells often contain a-subunits [253]. By electron microscopy these cells resemble densely granulated somatotrophs (Fig. 1.16a). They are fairly large, polyhedral cells with ovoid or slightly irregular nuclei. They have abundant electron-lucent cytoplasm which contains well-developed rough endoplasmic reticulum and a prominent Golgi apparatus which often harbors numerous immature secretory granules. The mitochondria vary from ovoid to rod-shaped; they have a light matrix and lamellar cristae. The distinctive feature of these cells is their unique population of secretory granules. Some are small, spherical, or slightly ovoid, electron-dense granules which measure 150–400 nm; they have tightly fitting limiting membranes. The larger granules are irregular, often extremely elongated structures which can measure from 350 to 2000 nm; they contain secretory material of variable electron density and are bound by a loosely fitting membrane. These secretory granules frequently show misplaced exocytosis with fusion of the limiting membrane to the cell membrane. Unlike the granule extrusions of lactotrophs, the contents that are emptied into the intercellular space persist and retain electron density. Ultrastructural

Chapter 1

(a)

Functional Anatomy of the Hypothalamic Pituitary Axis

25

(b)

FIGURE 1.16. (a) By electron microscopy, mammosomatotrophs resemble densely granulated somatotrophs but have numerous conspicuous large secretory granules of irregular shape and variable electron density, as well as the hallmark of prolactin secretion, the misplaced exocytosis (arrow). (Magnification ¥7300) (b) For the definitive evidence of bihormonal differentiation of mammosomatotrophs, ultrastructural immunocytology localizes both growth hormone (15 nm gold particles) and prolactin (40 nm gold particles) to the same cell, often within the same secretory granule. (Magnification ¥20160) (b) From Felix et al. [271]

immunocytology using the double immunogold technique documents the presence of both GH and PRL in a single cell, frequently within the same secretory granule (Fig. 1.16b). Hyperplasia of mammosomatotroph cells has been identified as a cause of gigantism of early onset [193] and has been associated with acromegaly and hyperprolactinemia in the McCune–Albright syndrome [190]. Although the pathogenesis of this lesion in human patients remains unclear, an animal model for this disorder is provided by giant mice transgenic for GHRH in whom chronic exposure to excess GHRH causes mammosomatotroph hyperplasia [194]; continued long-term GHRH stimulation leads to the development of mammosomatotroph adenoma in old animals [272]. In human patients, tumors composed of mammosomatotrophs are more common than assumed in the past; they are slowly growing adenomas most often found in young patients with gigantism and mild hyperprolactinemia [195,252]. Corticotrophs

A single cell type in the human pituitary is responsible for the production of the POMC molecule and its various derivatives, including ACTH, MSH, lipotropic hormone (LPH), and endorphins [195]. Corticotrophs comprise approximately 15–20% of the adenohypophysial cell popu-

lation. The vast majority of these cells are found in clusters in the central mucoid wedge of the adenohypophysis; occasional scattered cells are also found in the lateral wings of the anterior lobe. Corticotrophs are also the predominant cell type in the poorly developed intermediate lobe of the human pituitary where they are found scattered in follicular structures. By light microscopy these medium-sized cells have varying degrees of cytoplasmic basophilia and stain strongly with PAS; the affinity is attributed to the carbohydrate moiety present in ACTH precursors. They also stain with lead hematoxylin. The most reliable method of identifying corticotrophs is the immunoperoxidase technique which reveals strong granular cytoplasmic positivity for ACTH. The presence of a large unstained perinuclear lysosomal vacuole known as the “enigmatic body” may be helpful in identifying these cells. Corticotrophs can also be identified by in situ hybridization which localizes the POMC mRNA [273]. Immunocytologic studies have indicated that almost all corticotrophs contain immunoreactive MSH, LPH, endorphins and other fragments of the POMC molecule which are derived by differing posttranslational processing [158]. By electron microscopy (Fig. 1.17), corticotroph cells are oval or slightly angular medium-sized cells with spherical or oval eccentric nuclei and a spherical nucleolus which is

26

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Hypothalamic–Pituitary Function

FIGURE 1.18. In the pituitary of a patient treated with high doses of glucocorticoids, corticotrophs with Crooke’s hyaline change have glassy cytoplasm and their periodic acid–Schiff (PAS) positive secretory material is pushed to the rim of the cell; the large juxtanuclear globule (arrow head) is the enigmatic body. (PAS stain; magnification ¥256)

FIGURE 1.17. The nontumorous corticotroph has widely dispersed short rough endoplasmic reticulum membranes, a prominent Golgi complex (G) and numerous secretory granules of variable size, shape and electron density. The two highly characteristic features of this cell type are the perinuclear and cytoplasmic bundles of intermediate filaments (arrows) and the large lysosomal enigmatic body (EB). (Magnification ¥5150)

usually attached to the nuclear membrane [40]. The cytoplasm has varying electron density and is frequently moderately electron-opaque. The rough endoplasmic reticulum is moderately developed and takes the form of widely dispersed membranes. Numerous free ribosomes can be found. The Golgi apparatus is spherical or flattened and is often displaced by the “enigmatic body.” This large structure is membrane-bound and has an electron-dense periphery that exhibits acid phosphatase activity, confirming its lysosomal nature [274]. Mitochondria are spherical or oviod, with lamellar or tubular cristae and a moderately electron-dense matrix. Variable numbers of intermediate filaments of the cytokeratin type [275], previously described as type I microfilaments, are found in small bundles usually adjacent to the nucleus; they measure about 7 nm in width and show no periodicity. They vary considerably in amount and are not numerous under physiologic conditions. The secretory granules are usually numerous and extremely variable in size,

shape, and electron density. They may be spherical, flattened, dented, heart shaped, or tear-drop shaped. They vary in electron density and may be 150–700 nm in diameter, most measuring 150–400 nm. While secretory granules may be found lined up along the cell membrane, exocytosis is not described in this cell type. Immunoelectron microscopy identifies the various POMC-derived peptides in the secretory granules of corticotrophs; however, there is no evidence that the morphologic differences of granule populations reflect their content [38]. Corticotrophs are the first cell type to differentiate in the fetal pituitary [220,221]; however, they are found in significantly decreased numbers in anencephaly [276]; it has been suggested that after autonomous differentiation, they are dependent on hypothalamic factors for normal growth and development. Their numbers do not vary with age or changes in the hormonal environment; however, they do develop specific morphologic features that reflect changes in endocrine homeostasis. In the human pituitary exposed to glucocorticoid excess due to administration of exogenous corticosteroids or any cause of endogenous glucocorticoid hypersecretion (including ectopic secretion of ACTH), corticotrophs undergo a distinctive morphologic alteration known as Crooke’s hyaline change [195]. By light microscopy, the cells accumulate a glassy, homogeneous, slightly acidophilic substance in the cytoplasm; the PAS positivity and ACTH immunoreactivity, corresponding to secretory granules, are displaced to the perinuclear rim and the periphery of the cell [38] (Fig. 1.18). The hyaline material does not contain immunoreactive POMC derivatives. This material has been shown to be composed of keratin filaments and stains with several antibodies directed against low-molecular-weight keratin proteins [275]. By electron microscopy, Crooke’s hyaline

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27

FIGURE 1.20. Corticotrophs invade the neural lobe of the pituitary (N) in a normal age-related phenomenon that is unassociated with endocrinopathy. This case, the same as that illustrated in Fig. 1.18, shows that the population of corticotrophs participating in “basophil invasion,” like those lining the pars intermedia cysts (arrow head), do not undergo Crooke’s hyaline change. (Periodic acid–Schiff stain; magnification ¥102)

FIGURE 1.19. Corticotroph with Crooke’s change in the pituitary of a patient with cortisol excess has a ring of intermediate filaments filling the cytoplasm, surrounding the nucleus and trapping numerous secretory granules in a juxtanuclear location. (Magnification ¥5090)

material is composed of an accumulation of intermediate filaments, resembling the smaller bundles seen in nontumorous corticotrophs (Fig. 1.19). The accumulation of microfilaments can be so extensive as to occupy almost the entire cytoplasm, leaving only a small juxtanuclear Golgi region and a thin rim of secretory granules adjacent to the cell membrane. Crooke’s hyaline change is reversible [195]. In the pituitaries of patients with inadequately treated adrenal insufficiency, the corticotrophs become hypertrophied, with large nuclei, prominent nucleoli, and a poorly granulated cytoplasm. In cases of long-standing Addison’s disease, focal nodular hyperplasia of corticotrophs and corticotroph adenomas may be present [277]. The ultrastructural appearance of these “adrenalectomy” cells in the human pituitary has not been well documented. However, in adrenalectomized rats, the markedly enlarged cells contain abundant rough endoplasmic reticulum and Golgi membranes and an increased number of secretory granules [278]. Chronic administration of CRH to rats increases the number of ACTH-containing cells, but no significant ultra-

structural changes have been documented [279]. Morphometric analysis indicated that there was no change in cell area; the nuclear area increased slightly and the secretory granule diameter was greater in corticotrophs of CRHexposed animals. These data suggest that the proliferation of corticotrophs in Addison’s disease may be mediated by CRH, but the morphologic characteristics of adrenalectomy cells cannot be attributed solely to CRH excess. The pars intermedia corticotrophs also are strongly (PASpositive and exhibit intense immunostaining for ACTH and other POMC derivatives [195]. The border between the pars intermedia and the pars distalis is often indistinct; occasionally a thin layer of connective tissue is noticeable between these two portions of the adenohypophysis. On the posterior aspect of the pars intermedia, basophil cells are frequently identified, often in clusters within the neuropil of the pars nervosa. These clusters vary in number and in some cases they may be found spreading deeply into the neural lobe (Fig. 1.20). This process, known as “basophil invasion,” is found in older individuals; it is more frequent and pronounced with advancing age and is said to be more prominent in men than in women. Its functional significance is not known but it is not thought to be associated with endocrine abnormalities. Some studies have suggested that the presence of a-MSH immunoreactivity is indicative of intermediate lobe differentiation [280,281]. By electron microscopy, corticotrophs of the pars intermedia are smaller, more dense, and contain fewer intermediate filaments [192]. They are also known to be less sensitive to the feedback effect of glucocorticoids, in most cases showing absent or only mild filament accumulation in patients with glucocorticoid excess and lacking the Crooke’s hyaline change that is found in pars distalis corticotrophs of those patients [195].

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The corticotrophs of the pars intermedia and those which form the basophil invasion of the pars nervosa may give rise to a specific group of corticotroph adenomas. These tumors may represent one form of the silent corticotroph adenomas which are unassociated with signs or symptoms of ACTH excess [252]; they are probably capable of secreting biologically and immunologically active substances that remain uncharacterized. As is the case with pars intermedia corticotrophs, these tumors do not show striking accumulations of intermediate filaments and, in some cases, may lack filaments totally. The derivation of corticotroph cell adenomas from this subpopulation of corticotrophs is difficult to prove. The morphology of corticotrophs in most adenomas associated with ACTH excess resembles that of nontumorous corticotrophs [195,252]. Occasional tumors are composed of cells with Crooke’s hyaline change [252], suggesting that the adenomatous cells are sensitive to feedback inhibition by glucocorticoids. Morphologic studies of functioning corticotroph adenomas in vitro [282] confirm the response of most such tumors to CRH stimulation associated with slight ultrastructural changes indicative of increased hormone synthesis. Incubation of tumor cells with glucocorticoids inhibits ACTH release and causes accumulation of intermediate filaments; however, it remains to be seen if the threshold for suppression is altered in tumor cells.

idase technique reveals granular positivity for TSH; this stain reveals the characteristic angular shape of these cells and their long cytoplasmic processes that establish contact with the basement membrane (Fig. 1.21). Ultrastructurally, these cells are characterized by their marked angularity and well-developed cytoplasmic processes [40] (Fig. 1.22). The nucleus is spherical and often eccentric while the cytoplasm contains numerous short, slightly dilated rough endoplasmic reticulum profiles, a globoid Golgi complex and small, spherical secretory granules that measure 100–200 nm in diameter. When scanty, the secretory granules characteristically are aligned under the plasmalemma and may contain rod-shaped or irregularly electron-dense contents; more densely granulated cells have granules scattered throughout the cytoplasm. Although the numbers of thyrotrophs do not appear to vary with age, these cells undergo morphologic changes in association with altered hormonal status. In patients with primary hyperthyroidism, thyrotrophs are few and small;

Thyrotrophs

The least common cell type in the adenohypophysis is the thyrotroph. These comprise approximately 5% of the total adenohypophysial cell population and are found primarily singly or in small clusters in the anteromedial portion of the gland. These medium-sized cells are basophilic when stained with conventional dyes, contain PAS positivity, stain with aldehyde fuchsin and aldehyde thionin. The immunoperox-

FIGURE 1.21. Immunohistochemical localization of thyrotropin identifies scattered angular thyrotrophs in the nontumorous human autopsy pituitary. (Magnification ¥102)

FIGURE 1.22. Nontumorous thyrotrophs have welldeveloped cytoplasm with short, slightly dilated profiles of rough endoplasmic reticulum (arrow heads), a large Golgi region (G), small secretory granules lined up along the plasma membrane and small numerous secondary lysosomes (L). (Magnification ¥7440)

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29

subset of altered somatotrophs which are bihormonal [283]; the morphologic changes are reversible in the experimental animal. Gonadotrophs

FIGURE 1.23. In a patient with long-standing hypothyroidism, the cytoplasm of these “thyroidectomy cells” is dominated by the abundant widely dilated rough endoplasmic reticulum with flocculent contents; they also contain expanded Golgi complexes (G), few small secretory granules and prominent lysosomes (L). (Magnification ¥4380)

their ultrastructural features are not well documented. In cases of untreated primary hypothyroidism, thyrotrophs are released from the negative feedback effects of thyroid hormones and TSH secretion is increased. The number and size of thyrotrophs are increased; the enlarged cytoplasm is less strongly positive using the PAS, aldehyde fuchsin and aldehyde thionin stains, and TSH immunoreactivity is faint but diffuse. These cells exhibit strong nuclear positivity for Pit-1 [195,239]. By electron microscopy, these stimulated thyrotrophs, known as “thyroidectomy cells” or “thyroid deficiency cells,” have abundant dilated rough endoplasmic reticulum cisternae and an expanded Golgi complex, whereas the number of secretory granules is reduced [40] (Fig. 1.23). Large cytoplasmic spherical, PAS-positive lysosomal globules are prominent. With prolonged stimulation, nodular hyperplasia and thyrotroph adenomas may occur [195]; TRH may be implicated in the pathogenesis of this cell proliferation [252]. In the rat, there is evidence that at least some of the thyroidectomy cells may derive from a

The cells which produce the gonadotropins, FSH and LH are the gonadotrophs; these probably represent up to 10% of the human adenohypophysial cell population [195]. No reliable cell count is, however, available. In rats, the number of gonadotrophs containing each hormone varies with age, sex, and hormonal status [40]. Gonadotrophs are scattered throughout the pars distalis and comprise the major constituent of the pars tuberalis [284]. They stain with basic dyes, the PAS technique, aldehyde thionin, and aldehyde fuchsin. The immunoperoxidase technique reveals cytoplasmic positivity for FSH and LH, often both present in the same cell, indicating that one cell type is capable of producing both gonadotropins. There are, however, variations in the number of cells containing FSH and LH, indicating that some gonadotrophs contain only one of these two hormones and there are likely fluctuations with functional demand. Nuclear reactivity for SF-1 defines the gonadotroph population [237]. The ultrastructural features of gonadotrophs reveal a single cell population which is responsible for both FSH and LH production (Fig. 1.24). The large oval or elongated cells have spherical, eccentric nuclei which are oriented with distinct polarity at the base of the cell [40,252]. Their abundant, slightly dilated rough endoplasmic reticulum profiles contain a flocculent electron-lucent substance. The Golgi is prominent and globoid, composed of sacculi and vesicles associated with immature secretory granules. Mature secretory granules are unevenly distributed in the cytoplasm and fall into two groups which vary in size and density. The distribution of these two granule populations may show sexual dimorphism. In men, smaller secretory granules are usually numerous; they have an average diameter of 250 nm. In women, larger secretory granules predominate and measure 300–600 nm in diameter. Ultrastructural immunocytology localizes FSH and LH within the same cells and, in some cases, within the same secretory granule. Gonadotrophs are found in close proximity to the basement membrane and exhibit an intriguing intimate contact with lactotrophs. The latter frequently extend cell processes around gonadotrophs and there are intercellular junctions between the two cell types [285], suggesting paracrine interactions which are not well understood. Gonadotrophs are found in the pituitaries of fetuses and show sex-related dimorphism during gestation [220]. Between 15 and 25 weeks of gestation, pituitaries of female fetuses contain more numerous gonadotrophs than pituitaries of male fetuses. Throughout gestation, LH-containing cells predominate in male pituitaries, whereas the number of LHand FSH-containing cells are almost equal in females. This dimorphism correlates with differences in the levels of hypothalamic GnRH at the same stages of gestation. In the adult

30

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Hypothalamic–Pituitary Function

FIGURE 1.24. A nontumorous gonadotroph is large and oval with short profiles of dilated rough endoplasmic reticulum containing electron-lucent contents (*), ring-like Golgi complex (G) and large, dense but irregular secretory granules. (Magnification ¥8170)

pituitary, no such dimorphism of gonadotroph numbers has been identified; however, the ultrastructural differences in males and females have been described above. Gonadotroph adenomas of the adult pituitary show an even more striking fine-structural sexual dimorphism [195,286]. Gonadotroph adenomas in women are usually well-differentiated tumors consisting of polyhedral cells with long cytoplasmic processes, dilated profiles of well-developed endoplasmic reticulum, and the characteristic “honeycomb” Golgi apparatus, a large saccular structure associated with developing secretory granules. The secretory granules in this tumor type are spherical and membrane-bound with varying electron density; they measure about 400–450 nm in diameter. The tumors in adult men are less differentiated, with poorly to moderately developed rough endoplasmic reticulum profiles, a moderately developed Golgi complex and fewer, generally smaller secretory granules. Strong nuclear SF-1 positivity is a reliable marker [237]. Gonadotrophs of the pars distalis show morphologic changes which reflect their hormonal environment; in con-

trast, the gonadotrophs of the pars tuberalis show signs of functional inactivity in most circumstances. In patients treated with pharmacologic doses of estrogen, the gonadotroph cells are small and dense [195]. During pregnancy, the number of cells immunoreactive for FSH and/or LH is also significantly reduced [264]. Gonadotrophs are morphologically abnormal in Kallmann’s syndrome, an uncommon variant of hypothalamic hypogonadism due to GnRH deficiency and associated with anosmia [182]; there is a marked decrease in the number of these cells and the remaining small cells are only weakly positive for FSH and LH. Castration leads to characteristic morphology in gonadotrophs [40,195,287]. Prolonged lack of the negative feedback effect of gonadal steroids results in stimulation of gonadotrophs to secrete FSH and LH in higher quantities. These stimulated cells are enlarged and are found in increased numbers. They have vacuolated cytoplasm which may take the form of several small vacuoles or one large vacuole; the latter displaces the nucleus to the cell periphery, giving the cell a “signet ring” appearance. By electron microscopy, these so-called “gonadectomy cells,” “gonadal deficiency cells,” or “castration cells” (Fig. 1.25) have markedly dilated rough endoplasmic reticulum, which is responsible for the cytoplasmic vacuoles seen by light microscopy. The Golgi complex is enlarged and hormone storage is decreased but not lost, despite the marked increase in secretory activity, since secretory granules are present in reduced numbers. It has been suggested that secretory granule formation is bypassed and that discharge of hormone takes place in an unconventional way not discernible by electron microscopy. An unusual prominence of large active gonadotrophs has been noted in the nontumorous adenohypophysis of some women with PRL-producing adenomas [40]. The factors underlying this change are not clear, but it has been suggested as the explanation for the LH hyperresponse to GnRH which has been found in these patients. Follicular Cells

Follicular cells are found throughout the adenohypophysis surrounding follicles, lumina lined mainly by agranular or poorly granulated cells that are joined at their apex by junctional complexes [195,288]. By electron microscopy, it was shown that granulated adenohypophysial cells can form follicles around damaged cells which have disrupted cell membranes. The surrounding cells form specialized intercellular attachments and the participating cells undergo degranulation and dedifferentiation. These follicular cells may derive from somatotrophs, lactotrophs, or corticotrophs. The follicular cells form specialized junctions, macula adhaerens, between each other and with adjacent granulated adenohypophysial cells; in contrast, granulated adenohypophysial cells form only the less prominent zonulae adhaerentes. Follicles are found in areas with increased cell destruction, particu-

Chapter 1

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31

FIGURE 1.26. Immunolocalization of S-100 protein identifies a pituitary stellate cell with long branched cytoplasmic processes surrounding granulated adenohypophysial cells. (Magnification ¥256)

FIGURE 1.25. After castration, “gonadectomy cells” have markedly dilated rough endoplasmic reticulum and large globoid Golgi regions (G) which harbor forming secretory granules. (Magnification ¥4620)

been implicated as the source of FGF and vascular endothelial growth factor [293], but are not found in the pituitaries of anencephalics [294]. They are numerous in the compressed adenohypophysis at the periphery of adenomas. Some investigators have found no S-100-reactive sustentacular cells within pituitary adenomas [290]; in contrast, others have identified S-100-containing cells within tumors of several types [295]. They are also found in large numbers at the periphery of other pituitary lesions, such as abscesses, amyloid deposits and in the residual hypophysis after surgery, but not adjacent to metastatic tumor deposits, infarcts, or Rathke’s cleft cysts [296]. Null Cells

larly surrounding tumors [40]. They are assumed to play a role in isolating and processing cell debris. Some pituitary adenomas contain follicles. Folliculostellate Cells

Immunocytochemical studies have localized S-100 protein to a specific subtype of cells in the normal human pituitary gland [289,290]. These cells are agranular and do not contain immunoreactive hormones. Some also contain immunoreactivity for glial fibrillary acidic protein (GFAP). They have a characteristic morphology, with long, branched cytoplasmic processes embracing granulated adenohypophysial cells (Fig. 1.26). Because of confusion with the follicular cells described above, some authors have suggested that these cells be called “stellate cells” [289]. They are believed to have a supportive role similar to that played by the S-100-positive sustentacular cells of the adrenal medulla and carotid body. In addition, they are thought to play a role in paracrine regulation [291] and have been shown to produce IL-6, a cytokine which may participate in local regulation of hormone secretion [292]. These cells have also

The term null cell was used to identify cells that show evidence of adenohypophysial differentiation, possessing organelles required for hormone synthesis and storage, but whose differentiation is incomplete and does not resemble any of the known adenohypophysial cell types [297]. The term was applied initially to a group of tumors which possessed no histologic, immunocytochemical, or ultrastructural markers revealing their cellular differentiation and which were unassociated with clinical evidence of hormone excess. Application of increasingly sensitive techniques of cell identification have shown that most tumors with morphologic features of null cells are in fact gonadotroph adenomas [298,237]; however, only a small subpopulation of tumor cells may be hormonally active at any one time, as indicated by the reverse hemolytic plaque assay [299]. Immunohistochemical analyses [221] and studies of hormone mRNA [300] have shown that the majority of these tumors contain cells capable of producing glycoprotein hormone subunits. Focal ultrastructural differentiation along several lines has also been documented [301]. It has been suggested that these tumors may originate from uncommitted or

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committed stem cells or from dedifferentiated adenohypophysial cells. Null cells may also represent unstimulated or resting cells that are not actively engaged in hormone production and cannot be recognized with currently available investigative techniques [302]. The nontumorous pituitary contains cells that cannot be identified conclusively by ultrastructural criteria. These may represent resting cells, uncommitted or committed stem cells. In the fetal pituitary, a population of cells with features of the glycoprotein hormone cell line can be identified prior to the recognition of differentiated thyrotrophs or gonadotrophs [221]; a-subunit immunoreactivity is present in fetal glands at the same stage of gestation and b-subunits are not yet detected [220]. It has been suggested that these primitive cells which resemble null cells may represent the source of a-subunit and may be precursors of the glycoprotein hormone cell line; they may be the cell of origin of some null cell adenomas. The null cells are chromophobic and by immunocytochemistry, usually contain no hormones [195,252]. A small number of these cells may contain various pituitary hormones, most commonly a-subunit of the glycoprotein hormones followed by b-subunits of FSH and/or LH. They usually exhibit SF-1 nuclear staining, indicating gonadotrophin differentiation [237]. By electron microscopy, the cells are polyhedral with irregular nuclei. The poorly developed cytoplasm contains short, scattered rough endoplasmic reticulum profiles, an inconspicuous or only moderately developed Golgi apparatus and small, dense, rodshaped mitochondria. Secretory granules are few and small, measuring less than 250 nm in diameter, frequently with central electron-dense cores, peripheral electron-lucent halos, and a prominent limiting membrane (Fig. 1.27). Oncocytes

Oncocytes are large cells which have abundant acidophilic granular cytoplasm due to the numerous mitochondria that take up acid dyes [195,252]. By electron microscopy, they are characterized by a striking increase in the number and size of mitochondria. A varying number of oncocytic cells may be found in the normal human hypophysis [303]. The number of these cells appears to increase with advancing age, a phenomenon also found in other organs including thyroid, parathyroid, and salivary glands. Cells in hormonally active adenomas only rarely show oncocytic transformation; the tumors usually retain their fine structural and immunocytochemical markers and can be recognized as showing various degrees of oncocytic change. In contrast, null cells, which have no other identifying markers, more frequently undergo oncocytic change. Gonadotroph adenomas regularly contain a varying number of cells with oncocytic change, and the number of oncocytes in any given tumor often increases with tumor recurrence [195]. Oncocytomas are found more commonly in elderly patients. In vitro studies using tissue culture techniques [298] and the reverse hemolytic plaque

FIGURE 1.27. A null cell in the nontumorous human pituitary has poorly developed organelles in a scant cytoplasm. However, there are short profiles of rough endoplasmic reticulum (arrow heads) and a few small secretory granules, indicating the potential for hormone synthesis. This cell has no features which allow recognition of its differentiation. (Magnification ¥5950)

assay [299] have indicated that there is no functional difference between null cell adenomas and oncocytomas of the human pituitary. The factors underlying oncocytic change are unknown, but they may lie in abnormalities of the mitochondrial DNA which is independent of the cellular DNA [304]. Oncocytes are large polyhedral cells with abundant granular cytoplasm that varies from strongly acidophilic to chromophobic. These cells may show ill-defined purple coloration with trichrome stains using aniline blue; this may lead to misinterpretation as basophilia. The phosphotungstic acid–hematoxylin method is helpful to demonstrate mitochrondrial abundance. Oncocytes showing ultrastructural features of differentiated adenohypophysial cells may contain the appropriate immunoreactive hormone. The majority of oncocytes, which are thought to be derived from null cells of gonadotrophs contain no immunoreactive hormones or may contain a-subunit, FSH LH. They often have nuclear SF-1 reactivity [237].

Chapter 1

FIGURE 1.28. In the nontumorous human pituitary, an isolated oncocyte has abundant cytoplasm which is almost totally occupied by numerous spherulated mitochondria. Rough endoplasmic reticulum (arrow heads), a small Golgi complex (G) and small secretory granules suggest that the cell is capable of hormone synthesis and storage. (Magnification ¥5590)

By electron microscopy, oncocytes are readily identified as large, polyhedral cells with irregular nuclei and a cytoplasm almost totally filled with numerous mitochondria. Histologically some oncocytes are dark or acidophilic; these cells contain tightly packed oblong mitochondria with granular matrix and numerous transverse cristae. Other oncocytes may be histologically light, i.e., only slightly acidophilic or chromophobic; their crowded mitochondria appear swollen and rarefied, with loss of cristae. Oncocytes contain small, short profiles of rough endoplasmic reticulum, inconspicuous Golgi regions and few scattered secretory granules (Fig. 1.28). Others

Acidophil Stem Cells are recognized primarily in pituitary adenomas producing both GH and PRL [195,305]. They are rare and are not known to occur in the nontumorous pituitary. The cell type is assumed to derive from a common precursor of somatotrophs and lactotrophs; while

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mammosomatotrophs have been recognized as the source of GH and PRL early in gestation, no cells resembling tumorous acidophil stem cells have yet been identified in the human fetal pituitary [221]. Immunohistochemistry reveals the presence of PRL and GH in the cells. The positivity for PRL is usually stronger and patients with tumors composed of these cells have hyperprolactinemia; GH production may be reflected in the phenomenon known as “fugitive acromegaly.” By electron microscopy, these cells resemble sparsely granulated lactotrophs and have some features of somatotrophs. They are irregular in shape and have large nuclei containing fine chromatin and inconspicuous nucleoli. The cytoplasm contains scattered rough endoplasmic reticulum profiles and relatively small Golgi regions. The Golgi sacculi are flat and may contain a few forming granules. The mitochondria may be abundant, indicating oncocytic transformation, and often show focal cavitation as well as a unique mitochondrial gigantism not identified in other pituitary cell types. Giant mitochondria may reach the size of a nucleus and be visible by light microscopy as cytoplasmic vacuoles. The mitochondria retain their double limiting membranes but lose their cristae and are filled with electron-lucent granular matrix. Occasionally, they contain electron-dense tubular structures resembling tubules of centrioles and cilia. The secretory granules in this cell type are sparse and small, ranging from 50 to 300 nm. Characteristically, granule extrusions or “misplaced exocytoses” are identified, similar to those of sparsely granulated lactotrophs. In addition, these cells have a structural marker of sparsely granulated somatotrophs, the fibrous body, a juxtanuclear aggregate of smooth endoplasmic reticulum membranes, and keratin immunoreactive intermediate filaments. These features of lactotrophs and somatotrophs are frequently found in the same cell; ultrastructural immunocytochemistry confirms the localization of GH and prolactin in the same cell. A tumor type identified as silent subtype III adenoma is composed of chromophobic or slightly acidophilic cells which contain immunocytochemical evidence of multiple hormones. Staining for ACTH is generally weak, bendorphin immunoreactivity may be strong and the tumor cells may contain GH, PRL, and/or a-subunit immunoreactivity. In some cases, the tumors contain no recognizable hormones. By electron microscopy, large, elongated cells have an eccentric nucleus, a prominent nucleolus which may harbor spheridia, and a highly differentiated cytoplasm with abundant rough and smooth endoplasmic reticulum profiles and a well-developed Golgi apparatus. The secretory granules are small, measuring less than 200 nm in diameter, and have variable electron density; they are mainly spherical but occasionally irregular or dropshaped granules are found. These cells bear no similarity to any of the known nontumorous adenohypophysial cells and their cytodifferentiation is unknown. Other unclassified plurihormonal cells have been identified in the nontumorous pituitary and adenomas [40,195]. The

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most common types produce GH, a-subunit, TSH, and/or PRL in any combination. Some of these cells have ultrastructural features of somatotrophs or mammosomatotrophs, while others resemble thyrotrophs [40]. Studies of the rat pituitary have confirmed the existence of these cells and indicate a role for them in some pathologic states; for example, some somatotrophs transform into a population of thyroidectomy cells in hypothyroidism [283]. This common pattern of plurihormonality is easily explained by the expression of Pit-1, the transcription factor implicated in activation of the genes encoding GH, PRL and bTSH. However, occasionally other patterns of plurihormonality have been reported. Very rarely, cells producing POMC derivatives with ultrastructural features of corticotrophs may produce gonadotropins or other hormones both in human tumors [195,306] and in the rat nontumorous pituitary [307]. Conversely, cells with gonadotroph morphology may contain ACTH as well as gonadotropins [308]; it is intriguing, however, that ACTH, which is so commonly expressed in extrapituitary tumors, is rarely found in plurihormonal cells of the adenohypophysis.

Localization of Other Peptides in the Adenohypophysis A number of substances not initially thought to be of pituitary origin are detectable in adenohypophysial cells. Hypothalamic Adenohypophysiotropic Hormones

GHRH, somatostatin, TRH, and GnRH have been identified in pituitary adenohypophysial cells [308–311]. Large amounts of GHRH and somatostatin were measured in culture media of perifused normal pituitaries and GHsecreting adenomas and their release was stimulated by other peptides [309]. Preprosomatostatin mRNA has been detected in nontumorous adenohypophysis as well as several types of pituitary adenomas [310]. It is known that GnRH binds to gonadotrophs and becomes internalized [311]; however, it has also been shown that TRH and GnRH are released by pituitary cells maintained in culture for a prolonged period [308]. GnRH is colocalized with b-LH and some gonadotrophs also contain TRH. These data suggest production of hypothalamic peptides by de novo synthesis in a multipotential pituitary cell. Angiotensin II

Angiotensin II, as well as renin and angiotensin converting enzyme, have been localized in pituitary gonadotrophs [312]. These substances colocalize with b-LH. Angiotensin II has been shown to stimulate PRL release in vitro; this is a dosedependent phenomenon. It has also been found to stimulate ACTH and b-endorphin release. Lactotrophs and corticotrophs have high-affinity binding sites for angiotensin II. Bombesin/GRP

Bombesin-like immunoreactivity has been found in the pituitary gland [313]; however, cell localization has not been

demonstrated. Bombesin-related peptides stimulate pituitary hormone release, mediated by the hypothalamus. CGRP

CGRP is found in the pituitary in small amounts and the concentration is known to increase with age. It is unclear if this substance is present in adenohypophysial cells or in nerve terminals of the posterior lobe [314], since CGRP is found in the hypothalamus (see above). CCK

CCK is found in the adenohypophysis but in a form which differs from that present in other CCK-synthesizing tissues [315]. The pituitary is devoid of CCK-8 but contains substantial amounts of three large nonamidated pro-CCK fragments and small amounts of a-amidated CCK (CCK-58, CCK-33, component I). No immunocytochemical localization has been reported but CCK has been found in corticotroph adenomas in large amounts and in some tumors associated with acromegaly which also contained ACTH. It therefore seems that pituitary CCK derives from corticotrophs [316]. The form found in tumors is smaller and less sulfated than in the normal pituitary; tumors contain CCK-8-like forms similar to those found in brain and gut [315]. Galanin

In humans, galanin has been localized in corticotrophs of the nontumorous pituitary, including Crooke’s cells and cells of basophil invasion in the posterior lobe. Galanin is also present in some corticotroph adenomas [139,317]. In contrast, in rats, galanin mRNA is found in estrogen-induced pituitary tumors [318] and galanin has been localized in PRL- and GH-producing cells as well as some TSHcontaining cells [319]. The reason for this species-specific regulation is unknown. In the rat, galanin expression shows sexual dimorphism [319], is inducible by estrogen [318–320] and is altered by changes in adrenal, thyroid, or gonadal status [319]. It appears to be involved in the modulation of pituitary GH and PRL secretion, probably mediated by alterations in their hypothalamic regulation [140,141]. Gastrin

The neurohypophysis is known to contain gastrin [142], but there appears to be species-specific variation in the adenohypophysis. The human adenohypophysis contains only traces of this substance which is localized to corticotrophs [321] and corticotroph adenomas [322]. Analogy has been drawn to cosynthesis with POMC in other tissues; the significance of this colocalization is unknown and gastrin is not thought to act directly on the pituitary but rather in the hypothalamus. NPY

NPY immunoreactivity and mRNA are found in the pituitary and can be localized by immunocytochemistry

Chapter 1

in adenohypophysial cells consistent with a subset of thyrotrophs [323]. These cells increase in size and NPYimmunoreactivity after thyroidectomy. NPY may have autocrine or paracrine activity in the modulation of release of GH and gonadotropins. Neurotensin

Neurotensin has been localized to the anterior pituitary and is thought to be present in gonadotrophs. It is also known to be affected by thyroid status, being found in reduced amounts in both hypothyroidism and hyperthyroidism [151,324]. The role of neurotensin in the pituitary is not known. Substance P

Immunoreactive substance P has been found in the rodent anterior pituitary. In rats it is localized in gonadotrophs and lactotrophs [325], but in the guinea pig it is found in thyrotrophs [326]. Thyroidectomy increases and thyroid hormone treatment decreases the amount of substance P in the pituitary [152]; the amount also varies with levels of gonadal steriods. It is not clear if thyroid hormone and corticosteroids act directly to influence pituitary content. Substance P may play a local or paracrine role in the adenohypophysis. VIP

There is evidence that VIP is synthesized in the pituitary [327] in lactotrophs [127,328]; it is also inducible by estrogen and may be altered by other changes in the hormonal environment. Recently, it was found to be present in increased amounts in hypothyroidism [329] and VIP production by adenohypophysial cells in vitro was found to be stimulated by TRH as well as GHRH, but not by CRH or GnRH. Tissue culture studies have shown that VIP regulates tonic PRL secretion but does not affect TRH- or GHRHinduced PRL release [329]. Cytokines

Cytokines are known to modulate pituitary function and some have been shown to be produced in the hypothalamus (see above and references 161–166). IL-6 has been demonstrated in stellate cells within the pituitary [292] and is known to modulate GH and PRL responses to appropriate stimuli [164]. Growth Factors

Growth factors regulate cell replication and functional differentiation by directly altering the expression of specific genes. Several polypeptide growth factors regulate anterior pituitary hormone secretion. Growth factor action in the pituitary, however, must be understood in the context that the pituitary itself is a site of both synthesis and action of growth factors [330,331]. Several pituitary-derived growth factors are also regulated within the pituitary by peripheral hormones including triiodothyronine and hydrocortisone.

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Pituitary-derived growth factors include EGF, FGF, TGF-a and TGF-b, IGF, endothelin, and IL-1 [331]. Several partially characterized growth factors have also been described, including thyroid hormone-inducible growth factor, vascular endothelial growth factor mammary cell growth factor, chondrocyte growth factor, mammary cell growth factor, chondrocyte growth factor, and adipocyte growth factor [331]. EGF, a potent stimulator of PRL and ACTH secretion in vitro, also attenuates GH secretion [332–334]. EGF may also play a role in the regulation of LH secretion [335]. EGF has been localized in the pituitary within cells that produce glycoprotein hormones. EGF-receptor is expressed by normal human anterior pituitary and is upregulated in some adenomas, usually recurrent somatotroph tumors [336]. It has been speculated that EGF may mediate proliferation of thyrotrophs in hypothyroidism and of lactotrophs in the lactotroph hyperplasia of pregnancy [337]. TGF-a is expressed by pituitary cells where it has a membrane-anchored localization [338,339]; by immunohistochemistry it colocalizes with PRL and possibly with GH in the bovine pituitary [340]. TGF-a interacts with the EGF receptor and inhibits replication of rat pituitary tumor cells [341]. The regulation of lactotroph growth and differentiation by this substance suggests a potential role in promotion or facilitation of pituitary tumorigenesis [339] and in transgenic mice, targeting of TGF-a overexpression to lactotrophs results in lactotroph adenoma formation [343]. Activin, a member of the TGF-b gene family, suppresses GH secretion and somatotroph replication, and stimulates FSH secretion [343–346]. The inhibin subunits which comprise activin and inhibin and their mRNAs have been localized in pituitary gonadotrophs [309]. Activin-binding protein, which shows homology to follistatin, has been isolated in the rat pituitary and appears to block the action of activin [347]. Follistatin is expressed by normal gonadotrophs and is reduced or absent in gonadotroph adenomas, suggesting a role for this growth factor in the tumorigenic process [348]. The pituitary contains high concentrations of basic FGF [349]. There are several indications that the FGF family plays an important role in the regulation of pituitary function. It may be an important paracrine regulator of PRL secretion [170]; nonmitogenic concentrations of FGF stimulate TRHinduced PRL and TSH secretion in pituitary cells. Pituitaryderived FGF stimulates the replication of human pituitary cells in vitro [350]. FGF has also been implicated in the new arteriolar vessel formation reported to accompany prolactinomas [351]. FGF expression is upregulated in aggressive pituitary adenomas and this growth factor has been shown to be released by the adenoma cells [352], indicating its potential role in the control of adenohypophysial cell proliferation. Moreover, altered FGF receptors have been implicated in the dysregulated growth of pituitary tumors [353].

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Endothelin, a powerful vasoconstrictor, is expressed in the pituitary and stimulates FSH and LH secretion [354]. Pituitary-derived endothelin-3 is stimulated by IGF-I [355]. IGF-I mediates most of the peripheral growth promoting actions of GH [356]. GH-dependent IGF-I is secreted predominantly by the liver, and IGF-I participates in feedback regulation of GH gene transcription and secretion [357]. Regulation of pituitary IGF-I receptors also determines the action of the growth factor on somatotroph function [356]. IGF-I also stimulates the in vitro secretion of hypothalamic SRIH, further delineating its role in feedback regulation of GH secretion [358]. Pituitary expression of GH, IGF-I and IGF-I receptors have been documented [359,360] and may therefore mutually interact by a paracrine or autocrine feedback mechanism [361]. Regulation of endogenous or circulating pituitary growth factors and their respective receptors may be important paracrine or autocrine determinants of pituitary cell function and trophic hormone secretion. The role of these growth factors in the development of pituitary tumors is being intensively studied [232].

Classification of Adenohypophysial Pathology Many diseases can affect tissues in and adjacent to the sella turcica, giving rise to a wide spectrum of clinical pathology. The clinical problems may be manifest as pituitary hormone deficiency or pituitary hormone excess. In addition, associated problems arise from compression, invasion, injury, or destruction of other tissue components; symptoms such as visual field defects, nerve palsies and headaches are characteristic of mass effects on the hypothalamus and neurohypophysis. Adenohypophysial Hormone Deficiency

The clinical problem of pituitary hormone deficiency may be due to decreased secretion or to increased degradation of hormone. Characteristic adenohypophysial changes are often associated with abnormalities of hormone secretion. Hypopituitarism involving one, two, or more adenohypophysial hormones is usually attributed to either a decreased number of hormone-secreting cells or to reduced endocrine activity. Organic lesions such as massive necrosis, inflammatory, or neoplastic processes can replace large portions of the pituitary and account for reduction of hormone-secreting parenchyma [195]. Reduced endocrine activity can result from target gland hormone excess or administration of exogenous hormones feedback negatively to suppress hypophysial function. Alternatively, it may be due to decreased hypothalamic stimulation or interference with receptors which govern secretion of hormones. Various developmental abnormalities can reduce the amount of adenohypophysial parenchyma. Some of these have been reviewed above (see embryology). It is now recognized that mutations of PROP-1 or Pit-1 result in various patterns of pituitary hormone deficiency and hypoplasia

[229,230,233,245,247]. In pituitary aplasia, there is total absence of the adenohypophysis due to a defect in the formation of Rathke’s pouch; pituitary hypoplasia is a milder variant of this disorder. In anencephaly, the brain, including the hypophysiotropic hypothalamus, is missing; due to lack of neurohumoral control, the adenohypophysis is smaller than normal and there is marked reduction in the number of corticotrophs associated with a less striking reduction of other adenohypophysial cell types [276a]. Cornelia de Lange syndrome, a rare congenital cyst of Rathke’s pouch, may compress the adenohypophysis and lead to various degrees of hypopituitarism. Pituitary dystopia or ectopia rarely affects function. Tissue destruction leads to a decrease or absence of pituitary hormone-producing parenchyma. This may be caused by tumors, inflammatory or infiltrative processes, or vascular lesions. Symptoms of hypopituitarism develop only with reduction of more than 60% of anterior lobe tissue and is clinically significant when there is more than a 90% reduction of hormone-producing cells. Hormonally inactive tumors of the adenohypophysis represent approximately 25% of surgically removed pituitary adenomas [195]. The null cell adenomas and oncocytomas fall into this category; these cell types have been discussed above. Craniopharyngiomas are less common than pituitary adenomas [195]. They represent 3–5% of intracranial neoplasms and occur usually in children or young adults. They are more frequently suprasellar than intrasellar and can cause varying degrees of hypopituitarism due to compression or destruction of the adenohypophysis, pituitary stalk, or hypophysiotropic hypothalamus. Other primary neoplasms in and around the sella turcica include fibromas, angiomas, gliomas, meningiomas, granular cell tumors, paragangliomas, chordomas, and many others (see Chapter 15) [195]. These tumors produce no hormones. The majority are benign and may be discovered incidentally at postmortem examination. Occasionally they cause hypopituitarism and local symptoms. Primary sarcomas have been reported in patients who have previously received irradiation for the treatment of pituitary adenomas [363]. Metastatic malignancies are found in patients with widely disseminated neoplasms; reported incidences vary from 1 to 14% [47,362]; however, the true incidence of adenohypophysial involvement is not known, since small deposits are easily overlooked. Pituitary involvement is rarely manifest before patients are overwhelmed by the malignancy. Inflammatory conditions in the sella turcica may occur as acute processes, with purulent hypophysitis and abscess formation, due to severe septicemia or direct extension from adjacent tissues. Granulomatous inflammation may be seen in tuberculosis, syphilis, sarcoidosis, and giant-cell granuloma. The latter is an uncommon disorder of unknown etiology which may involve the pituitary gland selectively; it may represent localized sarcoidosis [364]. Lymphocytic hypophysitis is an autoimmune disorder which may cause

Chapter 1

extensive destruction of the adenohypophysis and subsequent hypopituitarism [365,366]. Vascular lesions such as hemorrhages may occur as complications of traumatic head injuries or in tumors. Infarction is seen focally as an incidental autopsy finding but significant lesions may be associated with diabetes mellitus, generalized carcinomas, cranio-cerebral trauma, increased intracranial pressure, infection, and in patients maintained on mechanical ventilators. Postpartum pituitary necrosis, known as Sheehan’s syndrome [367], occurs in pregnancy, during which time the pituitary is predisposed to vasospasm and infarction. When damage is significant, hypopituitarism is proportional to the degree of tissue destruction. Infiltrative disorders such as amyloidosis, hemochromatosis, and mucopolysaccharidosis, may impair adenohypophysial cell function. Hemochromatosis can give rise to selective hypogonadism due to preferential deposition of iron in pituitary gonadotrophs [368]. Feedback inhibition may cause cytologic alterations in the adenohypophysis. These changes have been described above. Selective deficiencies of adenohypophysial hormones may result from inherited genetic abnormalities or selective damage to one cell type. Isolated GH deficiencies are a genetically heterogeneous group of disorders that cause dwarfism. In most cases, there is a normal pituitary with normal numbers of granulated somatotrophs; these are attributed to GHRH deficiency or abnormality [172–175]. Selective deficiency of a specific pituitary transcription factor, Pit-1, causes selective congenital GH and PRL deficiency associated with TSH deficiency [369–371]; several animal models of pituitary dwarfism, the Snell, Jackson and Ames dwarf mice, have also been shown to have specific abnormalities of the Pit-1 gene [372]. Congenital isolated ACTH deficiency has been attributed to T pit mutations. Lymphocytic hypophysitis has been reported to cause selective destruction of corticotrophs; antibodies to other pituitary cell types are also detectable in patients with endocrine autoimmune diseases and may give rise to isolated pituitary hormone deficiency [365,366]. Short stature with features of isolated GH deficiency but elevated GH blood levels was found to be due to a deficiency of GH-binding protein, the extracellular domain of the GH receptor [373,374]. Adenohypophysial Hormone Excess

Excess hormone production by the pituitary is usually due to a pituitary tumor [195]. The most common lesion is adenoma; carcinoma occurs exceedingly rarely and may be associated with hormone excess. Hyperplasia and hypertrophy of hormone-secreting cells may result from increased stimulation by hypothalamic releasing hormones, or from loss of inhibition by hypothalamic-inhibiting hormones, or excess target organ hormones. The cytologic features of each adenoma type have been described above. The features of hyperplastic adenohypophysial cells have also been provided. Tumor-like lesions may also be associated with hypersecretion of adenohypophysial hormones. Lymphocytic

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hypophysitis may give rise to a mass lesion of the sella turcica associated with hyperprolactinemia [365,366]. The elevated PRL levels may be due to inflammation, injury of the stalk or hypothalamus or compression of those structures by the enlarged, inflamed gland. The disorder is frequently associated with pregnancy and it has been suggested that antibodies directed against hyperplastic lactotrophs may stimulate PRL release, interfere with dopamine receptors, or damage PRL cells causing release of stored PRL. Rarely, pituitary hormone excess has been associated with normal pituitary morphology [195]. The hyperfunction may be caused by intrinsic abnormalities of the cells which produce subtle changes that may not be detectable by conventional morphologic techniques.

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Functional Anatomy of the Hypothalamic Pituitary Axis

41

213 Cox BM, Baizman ER, Su T-P et al. Further studies on the nature and function of pituitary endorphins. Adv Biochem Psychopharmacol 1978;18: 183–189. 214 Ferrand R, Hraoui S. Origine exlusivement ectodermique de l’adénohypophyse chez la caille et le poulet; demonstration par la méthode des associations tissulaires interspécifiques. C R Soc Biol 1973;167:740–743. 215 Ferrand R, Pearse AGE, Polak JM, Le Douarin NM. Immunohistochemical studies on the development of avian embryo pituitary corticotrophs under normal and experimental conditions. Histochemistry 1974;38:133–141. 216 LeDouarin N. Particularités du noyau interphasique chez la caille japonaise (Coturnix coturnix japonica). Utilisation de ces particularités comme “marque biologique” dans les recherches sur les interactions tissulaires et les migrations cellulaires au cours de l’ontogénèse. Bull Biol Fr Belg 1969;103: 435–452. 217 Takor Takor T, Pearse AGE. Neuroectodermal origin of avian hypothalamohypophyseal complex: the role of the ventral neural ridge. J Embryol Exp Morphol 1975;34:311–325. 218 Daikoku S. Studies on the human foetal pituitary. 1. Quantitative observations. Tokushima J Exp Med 1958;5:200–213. 219 Baker BL, Jaffe RB. The genesis of cell types in the adenohypophysis of the human fetus as observed by immunocytochemistry. Am J Anat 1975;143: 137–162. 220 Asa SL, Kovacs K, Laszlo FA et al. Human fetal adenohypophysis. Histologic and immunocytochemical analysis. Neuroendocrinology 1986;43:308–316. 221 Asa SL, Kovacs K, Horvath E et al. Human fetal adenohypophysis. Electron microscopic and ultrastructural immunocytochemical analysis. Neuroendocrinology 1988;48:423–431. 222 Tremblay JJ, Lanctot C, Drouin J. The pan-pituitary activator of transcription, Ptxl (pituitary homeobox 1) acts in synergy with SF-1 and Pit-1 and is an upstream regulator of the Lim-homeodomain gene Lim3/Lhx3. Mol Endocrinol 1998;12:428–441. 223 Gage PJ, Camper SA. Pituitary Homeobox 2, a novel member of the bicoidrelated family of homeobox genes, is a potential regulator of anterior structure formation. Hum Mol Genet 1997;6:457–464. 224 Sheng HZ, Moriyama K, Yamashita T et al. Multistep control of pituitary organogenesis. Science 1997;278:1809–1812. 225 Bach I, Rhodes SJ, Pearse RV et al. P-Lim, a LIM homeodomain factor, is expressed during pituitary organ and cell commitment and synergizes with Pit-1. Proc Natl Acad Sci USA 1995;92:2720–2724. 226 Hermesz E, Machem S, Mahon KA. Rpx: a novel anterior-restricted homeobox gene progressively activated in the prechordal plate, anterior neural plate and Rathke’s pouch of the mouse embryo. Develop 1996;122:41–52. 227 Sornson MW, Wu W, Dasen JS et al. Pituitary lineage determination by the Prophet of Pit-1 homeodomain factor defective in Ames dwarfism. Nature 1996;384:327–333. 228 Gage PJ, Brinkmeier MLSLM, Knapp LT. The Ames dwarf gene, df, is required early in pituitary ontogeny for the extinction of Rpx transcription and initiation of lineage-specific cell proliferation. Mol Endocrinol 1996;10: 1570–1581. 229 Wu W, Cogan JD, Pfäffle RW et al. Mutations in PROP1 cause familial combined pituitary hormone deficiency. Nature Genet 1998;18:147–149. 230 Fofanova O, Takmura N, Kinoshita E. Compound heterozygous deletion of the prop-1 gene in children with combined pituitary hormone deficiency. J Clin Endocrinol Metab 1998;83:2601–2604. 231 Jackson SM, Barnhart KM, Mellon P et al. Helix-loop proteins are present and differentially expressed in different cell lines from the anterior pituitary. Mol Cell Endocrinol 1993;96:167–176. 232 Asa SL, Ezzat S. The cytogenesis and pathogenesis of pituitary adenomas. Endocr Rev 1998;19:798–827. 233 Asa SL, Ezzat S. Molecular determinants of pituitary cytodifferentiation. Pituitary 1999;1:159–168. 234 Lamolet B, Pulichino AM, Lamonerie T et al. A pituitary cell-restricted T box factor, T pit, activates POMC transcription in cooperation with Pitx homeoproteins. Cell 2001;104:849–859. 235 Poulin G, Turgeon B, Drouin J. NeuroD1/beta2 contributes to cell-specific transcription of the proopiomelanocortin gene. Mol Cell Biol 1997;17: 6673–6682. 236 Ingraham HA, Lala DS, Ikeda Y et al. The nuclear receptor steroidogenic factor 1 acts at multiple levels of the reproductive axis. Genes Dev 1994;8:2302–2312. 237 Asa SL, Bamberger A-M, Cao B et al. The transcription activator steroidogenic factor-1 is preferentially expressed in the human pituitary gonadotroph. J Clin Endocrinol Metab 1996;81:2165–2170.

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SECTION 1

Hypothalamic–Pituitary Function

238 Rosenfeld MG. POU-domain transcription factors: pou-er-ful developmental regulators. Genes Dev 1991;5:897–907. 239 Asa SL, Puy LA, Lew AM et al. Cell type-specific expression of the pituitary transcription activator Pit-1 in the human pituitary and pituitary adenomas. J Clin Endocrinol Metab 1993;77:1275–1280. 240 Friend KE, Chiou Y-K, Laws ER Jr et al. Pit-1 messenger ribonucleic acid is differentially expressed in human pituitary adenomas. J Clin Endocrinol Metab 1993;77:1281–1286. 241 Pellegrini I, Barlier A, Gunz G et al. Pit-1 gene expression in the human pituitary and pituitary adenomas. J Clin Endocrinol Metab 1994;79:189–196. 242 Day RN, Koike S, Sakai M et al. Both Pit-1 and the estrogen receptor are required for estrogen responsiveness of the rat prolactin gene. Mol Endocrinol 1990;4:1964–1971. 243 Drolet DW, Scully KM, Simmons DM et al. TEF, a transcription factor expressed specifically in the anterior pituitary during embryogenesis, defines a new class of leucine zipper proteins. Genes Dev 1991;5:1739–1753. 244 Li S, Crenshaw EB III, Rawson EJ et al. Dwarf locus mutants lacking three pituitary cell types result from mutations in the POU-domain gene pit-1. Nature 1990;347:528–533. 245 Tatsumi K, Miyai K, Notomi T et al. Cretinism with combined hormone deficiency caused by a mutation in the Pit-1 gene. Nature Genet 1992;1:56–58. 246 Pfäffle RW, DiMattia GE, Parks JS et al. Mutation of the POU-specific domain of Pit-1 and hypopituitarism without pituitary hypoplasia. Science 1992;257:1118–1121. 247 Radovick S, Nations M, Du Y et al. A mutation in the POU-homeodomain of Pit-1 responsible for combined pituitary hormone deficiency. Science 1992;257:1115–1118. 248 Puy LA, Asa SL. The ontogeny of pit-1 expression in the human fetal pituitary gland. Neuroendocrinology 1996;63:349–355. 249 Ikeda Y, Luo X, Abbud R et al. The nuclear receptor steroidogenic factor 1 is essential for the formation of the ventromedial hypothalamic nucleus. Mol Endocrinol 1995;9:478–486. 250 Yang H-J, Ozawa H, Kurosumi K. Ultrastructural changes in growth hormone cells in the rat anterior pituitary after thyroidectomy as studied by immunoelectron microscopy and enzyme nistochemistry. J Clin Electr Microsc 1989;22:269–283. 251 Ezzat S, Kontogeorgos G, Redelmeier DA et al. In vitro responsiveness of morphological variants of growth hormone-producing pituitary adenomas to octreotide. Eur J Endocrinol 1995;133:686–690. 252 Horvath E, Kovacs K. The adenohypophysis. In: Kovacs K, Asa SL, eds. Functional Endocrine Pathology 2nd ed. Boston: Blackwell Science, 1998; 247–281. 253 Sano T, Ohshima T, Yanad S. Expression of glycoprotein hormones and intracytoplasmic distribution of cytokeratin on growth hormone-producing pituitary adenomas. Pathol Res Pract 1991;187:530–533. 254 Neumann PE, Goldman JE, Horoupian DS, Hess MA. Fibrous bodies in growth hormone-secreting adenomas contain cytokeratin filaments. Arch Pathol Lab Med 1985;109:505–508. 255 Billestrup N, Swanson L, Vale W. Growth hormone-releasing factor stimulates proliferation of somatotrophs in vitro. Proc Natl Acad Sci USA 1986;83: 6854–6857. 256 Richardson SB, Twente S. Evidence that diminished pituitary responsitivity to GHRF is secondary to intracellular GH pool depletion. Am J Physiol 1988;254:E358–364. 257 Loras B, Yi JY, Durand A et al. GRF et adénomes somatotropes humains. Correlations in vivo et in vitro entre la libération de GH et les aspects morphologiques et immunocytochimiques. Ann Endocrinol 1985;46: 373–382. 258 Kawakita S, Asa SL, Kovacs K. Effects of growth hormone-releasing hormone (GHRH) on densely granulated somatotroph adenomas and sparsely granulated somatotroph adenomas in vitro: a morphological and functional investigation. J Endocrinol Invest 1989;12:443–448. 259 Beckers A, Stevenaert A, Kovacs K et al. The treatment of acromegaly with SMS 201–995. Advanc Biosci 1988;69:227–228. 260 Barkan AL, Lloyd RV, Chandler WF et al. Preoperative treatment of acromegaly with long-acting somatostatin analog SMS 201–995: shrinkage of invasive pituitary macroadenomas and improved surgical remission rate. J Clin Endocrinol Metab 1988;67:1040–1048. 261 Ezzat S, Snyder PJ, Young WF et al. Octreotide treatment of acromegaly. A randomized, multicenter study. Ann Intern Med 1992;117:711–718. 262 Asa SL, Felix I, Kovacs K, Ramyar L. Effects of somatostatin on somatotroph adenomas of the human pituitary: an in vitro functional and morphological study. Endocr Pathol 1990;1:228–235.

263 Baringa M, Bilezikjian LM, Vale WW, Rosenfeld MG, Evans RM. Independent effects of growth hormone releasing factor on growth hormone release and gene transcription. Nature 1985;314:279–281. 264 Herman V, Weiss M, Becker D, Melmed S. Hypothalamic hormonal regulation of human growth hormone gene expression in somatotroph adenoma cell cultures. Endocr Pathol 1990;1:236–244. 265a Zafar M, Ezzat S, Ramyar L et al. Cell specific expression of estrogen receptor in the human pituitary and its adenomas. J Clin Endocrinol Metab 1995;80:3621–3627. 265b Friend KE, Chiou TK, Lopes MBS et al. Estrogen receptor expression in human pituitary. J Clin Endocrinol Metab 1994;78:1497–1504. 266 Asa SL, Penz G, Kovacs K, Ezrin C. Prolactin cells in the human pituitary: a quantitative immunocytochemical analysis. Arch Pathol Lab Med 1982;106: 360–363. 267 Scheithauer BW, Sano T, Kovacs K et al. The pituitary gland in pregnancy: a clinicopathologic and immunohistochemical study of 69 cases. Mayo Clin Proc 1990;65:461–474. 268a Scheithauer BW, Kovacs K, Randall RV, Ryan N. Effects of estrogen on the human pituitary: a clinicopathologic study. Mayo Clin Proc 1989;64: 1077–1084. 268b Asa SL, Kelly MA, Grandy DK, Low MJ. Pituitary lactotroph adenomas develop after prolonged lactotroph hyperplasia in dopamine D2 receptordeficient mice. Endocrinology 1999;140:5348–5355. 269 Tindall GT, Kovacs K, Horvath E, Thorner MO. Human prolactin-producing adenomas and bromocriptine: a histological, immunocytochemical, ultrastructural, and morphometric study. J Clin Endocrinol Metab 1982;55: 1178–1183. 270 Esiri MM, Bevan JS, Burke CW, Adams CBI. Effect of bromocriptine treatment on the fibrous tissue component of prolactin-secreting and nonfunctioning macroadenomas of the pituitary gland. J Clin Endocrinol Metab 1986;63:383–388. 271 Felix IA, Horvath E, Kovacs K et al. Mammosomatotroph adenoma of the pituitary associated with gigantism and hyperprolactinemia. Acta Neuropathol 1986;71:76–82. 272 Asa SL, Kovacs K, Stefaneanu L et al. Pituitary adenomas in mice transgenic for growth hormone-releasing hormone. Endocrinology 1992;131:2083–2089. 273 Lewis ME, Sherman TG, Burke S et al. Detection of proopiomelanocortin mRNA by in situ hybridization with an oligonucleotide probe. Proc Natl Acad Sci USA 1986;83:5419–5423. 274 Horvath E, Ilse G, Kovacs K. Enigmatic bodies in human corticotroph cells. Acta Anat 1977;98:427–433. 275 Neumann PE, Horoupian DS, Goldman JE, Hess MA. Cytoplasmic filaments of Crooke’s hyaline change belong to the cytokeratin class. An immunocytochemical and ultrastructural study. Am J Pathol 1984;116:214–222. 276a Asa SL, Kovacs K. Functional morphology of the human fetal pituitary. Pathol Annu 1984;19(1):275–315. 276b Pilardzic D, Kovacs K, Asa SL. Pituitary morphology in human anencephalic fetuses. Neuroendocrinology 1997;65:164–172. 277 Scheithauer BW, Kovacs K, Randall RV. The pituitary gland in untreated Addison’s disease. A histologic and immunocytologic study of 18 adenohypophyses. Arch Pathol Lab Med 1983;107:484–487. 278 Siperstein ER, Allison VF. Fine structures of the cells responsible for secretion of adrenocorticotropin in the adrenalectomized rat. Endocrinology 1965;76: 70–79. 279 Asa SL, Kovacs K, Hammer GD et al. Pituitary corticotroph hyperplasia in rats implanted with a medullary thyroid carcinoma cell line transfected with a corticotropin-releasing hormone complementary deoxyribonucleic acid expression vector. Endocrinology 1992;131:715–720. 280 Coates PJ, Doniach I, Hale AC, Rees LH. The distribution of immunoreactive a-melanocyte-stimulating hormone cells in the adult human pituitary gland. J Endocrinol 1986;111:335–342. 281 McNicol AM. A study of intermediate lobe differentiation in the human pituitary gland. J Pathol 1986;150:169–173. 282 Horvath SE, Asa SL, Kovacs K et al. Human pituitary corticotroph adenomas in vitro: morphologic and functional reponses to corticotropin-releasing hormone and cortisol. Neuroendocrinology 1990;51:241–248. 283 Horvath E, Lloyd RV, Kovacs K. Propylthiouracyl-induced hypothyroidism results in reversible transdifferentiation of somatotrophs into thyroidectomy cells: a morphologic study of the rat pituitary including immunoelectron microscopy. Lab Invest 1990;63:511–520. 284 Asa SL, Kovacs K, Bilbao JM. The pars tuberalis of the human pituitary: a histologic immunohistochemical, ultrastructural and immunoelectron microscopic analysis. Virchows Archiv (Pathol Anat) 1983;399:49–59.

Chapter 1 285 Horvath E, Kovacs K. Gonadotroph adenomas of the human pituitary: sexrelated fine structural dichotomy. A histologic, immunocytochemical and electron microscopic study of 30 tumors. Am J Pathol 1984;117:429–440. 286 Denef C. Paracrine interactions in the anterior pituitary. Clin Endocrinol Metab 1986;15:1–32. 287 Kovacs K, Horvath E. Gonadotrophs following removal of the ovaries: a fine structural study of human pituitary glands. Endokrinologie 1975;66:1–8. 288 Horvath E, Kovacs K, Penz G, Ezrin C. Origin, possible function and fate of “follicular cells” in the anterior lobe of the human pituitary. Am J Pathol 1974;77:199–212. 289 Girod C, Trouillas J, Dubois MP. Immunocytochemical localization of S-100 protein in stellate cells (folliculo-stellate cells) of the anterior lobe of the normal human pituitary. Cell Tissue Res 1985;241:505–511. 290 Höfler H, Walter GF, Denk H. Immunohistochemistry of folliculo-stellate cells in normal human adenohypophyses and in pituitary adenomas. Acta Neuropathol 1984;65:35–40. 291 Baes M, Allaerts W, Denef C. Evidence for functional communication between folliculo-stellate cells and hormone-secreting cells in perifused anterior pituitary aggregates. Endocrinology 1987;120:685–691. 292 Vankelecom A, Carmeliet P, Van Damme J et al. Production of interleukin-6 by folliculo-stellate cells of the anterior pituitary gland in a histiotypic cell aggregate culture system. Neuroendocrinology 1989;49:102–106. 293 Ferrara N, Schweigerer L, Neufeld G et al. Pituitary follicular cells produce basic fibroblast growth factor. Proc Natl Acad Sci USA 1987;84: 5773–5777. 294 Coates PJ, Doniach I. Development of folliculo-stellate cells in the human pituitary. Acta Endocrinol 1988;119:16–20. 295 Lauriola L, Cocchia D, Sentinelli S et al. Immunohistochemical detection of folliculo-stellate cells in human pituitary adenomas. Virchows Archiv (Cell Pathol) 1984;47:189–197. 296 Nishioka H, Llena JF, Hirano A. Immunohistochemical study of folliculostellate cells in pituitary lesions. Endocr Pathol 1991;2:155–160. 297 Kovacs K, Horvath E, Ryan N, Ezrin C. Null cell adenomas of the human pituitary. Virchows Archiv (Pathol Anat) 1980;387:165–174. 298 Asa SL, Cheng Z, Ramyar L et al. Human pituitary null cell adenomas and oncocytomas in vitro: effects of adenohypophysiotropic hormones and gonadal steroids on hormone secretion and tumor cell morphology. J Clin Endocrinol Metab 1992;74:1128–1134. 299 Yamada S, Asa SL, Kovacs K et al. Analysis of hormone secretion by clinically nonfunctioning human pituitary adenomas using the reverse hemolytic plaque assay. J Clin Endocrinol Metab 1989;68:73–80. 300 Jameson JL, Klibanski A, Black PMcL et al. Glycoprotein hormone genes are expressed in clinically nonfunctioning pituitary adenomas. J Clin Invest 1987;80:1472–1478. 301 Kontogeorgos G, Horvath E, Kovacs K et al. Null cell adenoma of the pituitary with features of plurihormonality and plurimorphous differentiation. Arch Pathol Lab Med 1991;115:61–64. 302 Kovacs K, Asa SL, Horvath E et al. Null cell adenomas of the pituitary: attempts to resolve their cytogenesis. In: Lechago J, Kameya T, eds. Endocrine Pathology Update. Philadelphia: Field and Wood, 1990;17–31. 303 Kovacs K, Horvath E, Bilbao JM. Oncocytes in the anterior lobe of the human pituitary gland. A light and electron microscopic study. Acta Neuropathol 1974;27:43–54. 304 Carafoli E. Mitochondrial pathology: an overview. Ann NY Acad Sci 1986; 488:1–18. 305 Horvath E, Kovacs K, Singer W et al. Acidophil stem cell adenoma of the human pituitary: clinicopathological analysis of 15 cases. Cancer 1981;47: 761–771. 306 Sano T, Kovacs K, Asa SL, Smyth HS. Immunoreactive luteinizing hormone in functioning corticotroph adenomas of the pituitary. Immunohistochemical and tissue culture studies of two cases. Virchows Archiv (Pathol Anat) 1990;471: 361–367. 307 Moriarty GC, Garner LL. Immunocytochemical studies of cells in the rat adenohypophysis containing both ACTH and FSH. Nature 1977;265:356–358. 308 May V, Wilber JF, U’Prichard DC, Childs GV. Persistence of immunoreactive TRH and GnRH in long-term primary anterior pituitary cultures. Peptides 1987;8:543–558. 309 Joubert (Bression) D, Benlot C, Lagoguey A et al. Normal and growth hormone (GH)-secreting adenomatous pituitaries release somatostatin and GH-releasing hormone. J Clin Endocrinol Metab 1989;68:572–577. 310 Pagesy P, Li JY, Rentier-Delrue F et al. Evidence of pre-prosomatostatin mRNA in human normal and tumoral anterior pituitary gland. Mol Endocrinol 1989;3:1289–1294.

Functional Anatomy of the Hypothalamic Pituitary Axis

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311 Duello TM, Nett TM, Farquhar MG. Fate of a gonadotropin releasing hormone agonist internalized by rat pituitary gonadotrophs. Endocrinology 1983;112:1–10. 312 Deschepper CF, Crumrine DA, Ganong WF. Evidence that the gonadotrophs are the likely site of production of angiotensin II in the anterior pituitary of the rat. Endocrinology 1986;119:36–43. 313 Major J, Ghatei MA, Bloom SR. Bombesin-like immunoreactivity in the pituitary gland. Experientia 1983;39:1158–1159. 314 Zaidi M, Abeyasekera G, MacIntyre I. Calcitonin gene-related peptide: endocrine distribution and characterization of circulating forms. J Endocrinol Invest 1989;12:699–704. 315 Rehfeld JF. Preprocholecystokinin processing in the normal human anterior pituitary. Proc Natl Acad Sci USA 1987;84:3019–3023. 316 Rehfeld JF, Lindholm J, Andersen BN et al. Pituitary tumors containing cholecystokinin. N Engl J Med 1987;316:1244–1247. 317 Vrontakis ME, Sano T, Kovacs K, Friesen HG. Presence of galanin-like immunoreactivity in nontumorous corticotrophs and corticotroph adenomas of the human pituitary. J Clin Endocrinol Metab 1990;70:747–751. 318 Vrontakis ME, Peden LM, Duckorth MC, Friesen HG. Isolation and characterization of a complementary DNA (galanin) clone from estrogeninduced pituitary tumor messenger RNA. J Biol Chem 1987;262: 16755–16758. 319 O’Halloran DJ, Jones PM, Steel JH et al. Effect of endocrine manipulation on anterior pituitary galanin in the rat. Endocrinology 1990;127: 467–475. 320 Kaplan LM. Gabriel SM, Koenig JI et al. Galanin is an estrogen-inducible, secretory product of the rat anterior pituitary. Proc Natl Acad Sci USA 1988;85:7408–7412. 321 Larson LI, Rehfeld JF. Pituitary gastrins occur in corticotrophs and melanotrophs. Science 1981;213:768–770. 322 Bardram L, Lindholm J, Rehfeld JF. Gastrin in pituitary tumors. Acta Endocrinol 1987;115:419–422. 323 Jones PM, Ghatei MA, Steel J et al. Evidence for neuropeptide-Y synthesis in the rat anterior pituitary and the influence of thyroid hormone status: comparison with vasoactive intestinal peptide, substance P, and neurotensin. Endocrinology 1989;125:334–341. 324 Goedert M, Lightman SL, Nagy IT et al. Neurotensin in the rat anterior pituitary gland. Nature 1982;298:163–165. 325 Morel G, Chayvialle JA, Kerdelhue B, Dubois PM. Ultrastructural evidence for endogenous substance P-like immunoreactivity in the rat pituitary gland. Neuroendocrinology 1982;35:86–92. 326 de Palatis LR, Fiorindo RP, Ho RH. Substance P immunoreactivity in the anterior pituitary gland of the guinea pig. Endocrinology 1982;110: 282–284. 327 Arnaout MA, Garthwaite TL, Martinson DR, Hagen TC. Vasoactive intestinal polypeptide is synthesized in anterior pituitary tissue. Endocrinology 1986;119: 2052–2057. 328 Steel JH, Gon G, O’Halloran DJ et al. Galanin and vasoactive intestinal polypeptide are colocalised with classical pituitary hormones and show plasticity of expression. Histochemistry 1989;93:183–189. 329 Lam SK, Reichlin S. Pituitary vasoactive intestinal peptide regulates prolactin secretion in the hypothyroid rat. Neuroendocrinology 1989;50: 524–528. 330 Webster J, Ham J, Bevan JS, Scanlon MF. Growth factors and pituitary tumors. Trends Endocrinol Metab 1989;1:95–98. 331 Ezzat S, Melmed S. The role of growth factors in the pituitary. J Endocrinol Invest 1990;13:691–698. 332 White BA, Bancroft FC. Epidermal growth factor and thyrotropin-releasing hormone interact synergistically with calcium to regulate prolactin mRNA levels. J Biol Chem 1983;258:4618–4622. 333 Murdoch GH, Potter E, Nicolaisen AK et al. Epidermal growth factor rapidly stimulates prolactin gene transcription. Nature 1982;300:192–194. 334 Polk DH, Ervin MG, Padbury JF et al. Epidermal growth factor acts as a corticotropin-releasing factor in chronically catheterized fetal lambs. J Clin Invest 1987;79:984–988. 335 Miyake A, Tasaka K, Otsuka S et al. Epidermal growth factor stimulates secretion of rat pituitary luteinizing hormone in vitro. Acta Endocrinol 1985;108:175–178. 336 Leriche VK, Asa SL, Ezzat S. Epidermal growth factor and its receptor (EGFR) in human pituitary adenomas: EGF-R correlates with tumor aggressiveness. J Clin Endocrinol Metab 1996;81:656–662. 337 Kasselberg AG, Orth DN, Gray ME, Stahlman MT. Immunocytochemical localization of human epidermal growth factor/urogastrone in several human tissues. J Histochem Cytochem 1985;33:315–322.

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338 Mueller SG, Kobrin MS, Paterson AJ, Kudlow JE. Transforming growth factor-a expression in the anterior pituitary gland: regulation by epidermal growth factor and phorbol ester in dispersed cells. Mol Endocrinol 1989;3:976–983. 339 Ezzat S, Walpola IA, Ramyar L et al. Membrane-anchored expression of transforming growth factor-≤ in human pituitary adenoma cells. J Clin Endocrinol Metab 1995;80:534–539. 340 Kobrin MS, Asa SL, Samoondar J, Kudlow JE. a-Transforming growth factor in the bovine anterior pituitary gland: secretion by dispersed cells and immunohistochemical localization. Endocrinology 1987;121:1412–1416. 341 Ramsdell JS. Transforming growth factor-a and -b are potent and effective inhibitors of GH4 pituitary tumor cell proliferation. Endocrinology 1991;128: 1981–1990. 342 McAndrew J, Paterson AJ, Asa SL et al. Targeting of transforming growth factor-a expression to pituitary lactotrophs in transgenic mice results in selective lactotroph proliferation and adenomas. Endocrinology 1995;136: 4479–4488. 343 Ying S-Y. Inhibins, activins, and follistatins: gonadal proteins modulating the secretion of follicle-stimulating hormone. Endocr Rev 1988;9:267–293. 344 Roberts V, Meunier H, Vaughan J et al. Production and regulation of inhibin subunits in pituitary gonadotropes. Endocrinology 1989;124:552–554. 345 Billestrup N, González-Manchón C, Potter E, Vale W. Inhibition of somatotroph growth and growth hormone biosynthesis by activin in vitro. Mol Endocrinol 1990;4:356–362. 346 Corrigan AZ, Bilezikjian LM, Carroll RS et al. Evidence for an autocrine role of activin B within rat anterior pituitary cultures. Endocrinology 1991;128:1682–1684. 347 Kogawa K, Nakamura T, Sugino K et al. Activin-binding protein is present in pituitary. Endocrinology 1991;128:1434–1440. 348 Penabad JL, Bashey HM, Asa SL et al. Decreased follistatin gene expression in gonadotroph adenomas. J Clin Endocrinol Metab 1996;81:3397–3403. 349 Gospodarowicz D, Ferrara N, Schweigener L, Neufeld G. Structural characterization and biological functions of fibroblast growth factor. Endocr Rev 1987;8:95–114. 350 Prysor-Jones RA, Silverlight JJ, Jenkins JS. Oestradiol, vasoactive intestinal peptide and fibroblast growth factor in the growth of human pituitary tumour cells in vitro. J Endocrinol 1989;120:171–177. 351 Elias KA, Weiner RI. Direct arterial vascularization of estrogen-induced prolactin-secreting anterior pituitary tumors. Proc Natl Acad Sci USA 1984;81:4549–4553. 352 Ezzat S, Smyth HS, Ramyar L, Asa SL. Heterogeneous in vivo and in vitro expression of basic fibroblast growth factor by human pituitary adenomas. J Clin Endocrinol Metab 1995;80:878–884. 353 Ezzat S, Zheng L, Zhu XF, Wu GE, Asa SL. Targeted expression of a human pituitary tumor-derived isoform of FGF receptor-4 recapitulates pituitary tumorigenesis. J Clin Invest 2002;109:69–78. 354 Stojlkovic SS, Merelli F, Iida T et al. Endothelin stimulation of cystolic calcium and gonadotropin secretion in anterior pituitary cells. Science 1990;248:1663–1666. 355 Matsumoto H, Suzuki N, Shiota K et al. Insulin-like growth factor-I stimulates endothelin-3 secretion from rat anterior pituitary cells in primary culture. Biochem Biophys Res Commun 1990;172:661–668.

356 Prager D, Melmed S, Fagin J. Feedback regulation of growth hormone gene expression by insulin-like growth factor I. In: LeRoith D, Raizada MK, eds. Molecular and Cellular Biology of Insulin-like Growth Factors and Their Receptors. New York: Plenum Press, 1989:57–71. 357 Yamashita S, Melmed S. Insulin-like growth factor I regulation of growth hormone gene transcription in primary rat pituitary cells. J Clin Invest 1987;79:449–452. 358 Berelowitz M, Szabo M, Frohman LA et al. Somatomedin-C mediates growth hormone negative feedback by effects on both the hypothalamus and the pituitary. Science 1982;212:1279. 359 Han VKM, Lund PK, Lee DC, D’Ercole AJ. Expression of somatomedin/ insulin-like growth factor messenger ribonucleic acids in the human fetus: identification, characterization, and tissue distribution. J Clin Endocrinol Metab 1998;66:422–429. 360 Yamashita S, Weiss M, Melmed S. Insulin-like growth factor I regulates GH secretion and mRNA levels in human pituitary tumor cells. J Clin Endocrinol Metab 1986;63:730–735. 361 Asa SL, Coschigano KT, Bellush L et al. Evidence for growth hormone (GH) autoregulation in pituitary somatotrophs in GH antagonist-transgenic and GH receptor-deficient mice. Am J Pathol 2000;156:1009–1015. 362 Asa SL, Kovacs K. Histological classification of pituitary disease. J Clin Endocrinol Metab 1983;12:567–596. 363 Shi T, Farrel MA, Kaufmann JCL. Fibrosarcoma Complicating irradiated pituitary adenoma. Surg Neurol 1984;22:277–283. 364 Del Pozo JM, Roda JE, Montoya JG et al. Intrasellar granuloma. Case Report. J Neurosurg 1980;53:717–719. 365 Asa SL, Bilbao JM, Kovacs K et al. Lymphocytic hypophysitis of pregnancy resulting in hypopituitarism. A distinct clinicopathologic entity. Ann Intern Med 1981;95:166–171. 366 Thodou E, Asa SL, Kontogeorgos G et al. Lymphocytic hypophysitis: Clinicopathologic findings. J Clin Endocrinol Metab 1995;80:2302–2311. 367 Sheehan HL, Stanfield JR. The pathogenesis of post-partum necrosis of the anterior lobe of the pituitary gland. Acta Endocrinol 1961;37:479–510. 368 Bergeron C, Kovacs K. Pituitary siderosis. A histologic and immunocytologic study. Arch Intern Med 1979;139:248-249. 369 Pfäffle RW, DiMattia GE, Parks JS et al. Mutation of the POU-specific domain of Pit-1 and hypopituitarism without pituitary hypoplasia. Science 1992;257:1118–1121. 370 Radovick S, Nations M, Du Y et al. A mutation in the POU-homeodomain of Pit-1 responsible for combined pituitary hormone deficiency. Science 1992;257:1115–1118. 371 Tatsumi K, Miyai K, Notomi T et al. Cretinism with a combined hormone deficiency caused by a mutation in the Pit-1 gene. Nature Gen 1992;1:56–58. 372 Li S, Crenshaw EB III, Rawson EJ et al. Dwarf locus mutants lacking three pituitary cell types result from mutations in the POU-domain gene pit-1. Nature 1990;347:528–533. 373 Daughaday WH, Trivedi B. Absence of serum growth hormone binding protein in patients with growth hormone receptor deficiency (Laron dwarfism). Proc Natl Acad Sci USA 1987;84:4636–4640. 374 Baumann G, Shaw MA, Winter RJ. Absence of the plasma growth hormonebinding protein in Laron-type dwarfism. J Clin Endocrinol Metab 1987;65: 814–816.

C h a p t e r

2 Adrenocorticotropin Mark A. Herman Joseph A. Majzoub

The anterior pituitary forms via invagination of the pharyngeal stomodeum in the region of contact with the diencephalon. By week 5 of human gestation, this invagination, termed Rathke’s diverticulum, has formed, but the downward migration of the diencephalic diverticulum, destined to be the neurophypophysis, has not yet commenced. It is at this time that adrenocorticotropin (ACTH) is first detectable by immunocytochemistry in the collection of cells within Rathke’s diverticulum which are furthest from contact with the diencephalon [1]. By 8 weeks of gestation, ACTH is detectable by radioimmunoassay of both fetal pituitary tissue and fetal blood [1]. The hypophyseal–portal vascular system forms between 8 and 14 weeks gestation, dating the earliest time after which hypothalamic corticotropin releasing factors may function to regulate fetal pituitary ACTH [1]. By 14 weeks gestation, release of ACTH from human fetal corticotrophs is highly responsive to exposure to corticotropin releasing hormone (CRH) in vitro [2]. The intensity of immunohistochemical ACTH staining in the anterior pituitary increases progressively from the mid-first through the end of the second trimester. In contrast, it is only after 21 weeks gestation that ACTH-positive cells begin to appear in the pars intermedia of the human fetal pituitary, defined as that region between Rathke’s cleft anteriorly and the neurohypophysis posteriorly. The ACTHcontaining cells in this region are epithelial-like, in contrast to the more angular, ovoid appearance of corticotrophs in the anterior pituitary [1].

those containing a-L-fucose and complex N-glyosylprotein, as well as terminal b-galactose. Corticotrophs constitute between 10 and 20% of the cell population of the anterior pituitary [3]. They are large cells with an ovoid or angular shape. They occur either singly or in clumps. They are most numerous in the midsagittal region of the pituitary (median wedge) but also occur in the lateral wings of the gland. Although the adult human pituitary does not contain an intermediate lobe, the junctional zone between the anterior and posterior lobes is known as the zona intermedia. This region, derived from the portion of Rathke’s diverticulum posterior to Rathke’s cleft, contains scattered cells immunopositive for ACTH. Some of these ACTHcontaining cells appear to extend into the posterior pituitary, a feature known as basophilic invasion [3]. These areas of apparent migration of corticotrophs from the zona intermedia into the posterior pituitary occur focally along the border between these two regions. In humans, in addition to the sellar pituitary, a pharyngeal pituitary exists which is located in either the sphenoid sinus or within the sphenoid bone. It consists of pituitarylike tissue approximately 2 to 5 mm by 0.2 mm in size. It is connected to the sellar gland by transsphenoidal vessels [4]. Only 1% to 2% of the cells in the pharyngeal pituitary contain immunoreactive ACTH, in contrast to approximately 14% of the cells in the sellar pituitary [4]. The pharyngeal pituitary is thought to arise as a rest of tissue resulting from the normal migration of cells from Rathke’s pouch to the sella turcica. There have been several reports of Cushing’s disease due to corticotroph adenomas arising in the pharyngeal pituitary [4,5].

Adult Anatomy

Molecular Embryology

Corticotrophs were initially identified by their basophilic staining. This has been subsequently found to be due to the presence of complex sugars in corticotrophs, principally

Tremendous progress has been made in recent years concerning the molecular mechanisms controlling pituitary organogenesis and pituitary cell-lineage specification in

CELLS OF ORIGIN

Fetal Anatomy

45

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animal models. A complex network of transcriptional events mediated by extrinsic and intrinsic signals has been implicated in the determination and a stereotypical spatiotemporal differentiation of the five trophic cell types of the mature anterior pituitary gland. Intrinsic Signals

RPX (for Rathke’s pouch homeobox), also known as HESX1 (for homeobox gene expression in embryonic stem cells) is the earliest known marker for the pituitary primordium [6]. HESX1/RPX is a member of the paired-like class of homeobox genes. Early HESX1/RPX expression occurs in the anterior midline endoderm and prechordal plate precursor, followed by expression in the cephalic neural plate [6]. Expression becomes progressively restricted anteriorly in the cephalic neural plate in a distribution consistent with tissue known by fate mapping studies to form the primordium of the anterior pituitary [7–10]. HESX1/RPX expression ultimately becomes restricted to Rathke’s pouch, and down regulation of HESX1/RPX in the pouch coincides with the differentiation of pituitary-specific cell types [6]. Embryonic mice lacking Rpx demonstrate variable abnormalities of hypothalamic and pituitary morphogenesis, reduced prosencephalon, anophthalmia or microophthalmia, and defective olfactory development. Neonatal mice have abnormalities in the corpus callosum, the anterior and hippocampal commisures, and the septum pellucidum [11]. These abnormalities are reminiscent of defects seen in a heterogeneous group of human disorders known as septooptic dysplasia (SOD). Deficits in SOD include optic nerve hypoplasia, abnormalities of the midline brain structures, and abnormalities of the hypothalamic-pituitary axis. Patients commonly present with endocrinopathies including hypoglycemia and adrenal crisis, which may signal growth hormone, ACTH, or thryoid-stimulating hormone deficiency [12]. Dattani and colleagues have identified two siblings with SOD who are homozygous for missense mutations within the HESX1 homeodomain which prevents it from binding target DNA [11]. The siblings identified with HESX1 mutations presented with hypoglycemia hours after birth and demonstrated hypopituitarism, substantiating a role for HESX1 in human pituitary development [11,13]. The corticotroph lineage appears to diverge relatively early from the other cell types of the anterior pituitary. LHX3, a LIM-type homeodomain protein, is essential for growth of Rathke’s pouch and determination of pituitary cell lineages [14]. In Lhx3-deficient mice, proopiomelanocortin (Pomc) was detected in a small cohort of cells at the ventral base of the pouch remnant which roughly corresponds to the position of the first presumptive corticotroph cells to differentiate in the wild-type pouch [14]. Although some pouch cells were able to differentiate and express Pomc, these cells failed to proliferate. An insufficient or non-functional corticotroph cell mass may have caused the hypoplastic adrenal cortices noted in these animals [14].

Ptx1 is a bicoid-related homeobox transcription factor identified based upon its ability to activate transcription of Pomc in the corticotroph derived AtT-20 cell line [15] and its ability to interact with the transactivation domain of Pit-1 [16]. It is expressed in the primordial Rathke’s pouch, oral epithelium, first branchial arch, the duodenum, and hindlimb [16]. Ptx1 is expressed in all mature pituitary cells, but is differentially expressed during pituitary development in different cell types [17]. In the mature pituitary, the highest levels of expression are found within corticotrophs [15]. In addition to activating transcription of Pomc, Ptx1 is required for sustained expression of Lhx3 [18], directly activates transcription of the common a-glycoprotein subunit (a-GSU) [16], synergizes with Pit-1 on the growth hormone and prolactin promoter [18], and synergizes with SF-1 on the promoter of the lutenizing hormone b (LH-b) gene [18]. Ptx2, also known as RIEG, is an additional bicoid-related homeobox gene 97% identical to Ptx1 [19]. Ptx2 is also differentially expressed in the pituitary and is excluded from corticotrophs [20], suggesting that some of the functions ascribed to Ptx1 may be performed by Ptx2. Expression of Pit-1, a POU domain transcription factor, is restricted to the anterior pituitary and required for the development of thyrotophs, lactotrophs, and somatotrophs [21]. Consistent with this, patients with mutations in PIT-1 have normal hypothalamic-pituitary-advanced (HPA) axis function [22]. Prophet of PIT-1 (PROP-1) is required for PIT-1 expression, and mutation of its gene is also associated with deficiencies in the development of thyrotrophs, lactotrophs, and somatotrophs in the Ames dwarf mouse and in humans. In a small number of patients with mutations in PROP1, modest impairment of ACTH secretion has been reported [23]. Developmentally essential, cell line-restricted factors have been identified for a number of different pituitary cell types. SF-1, an orphan nuclear receptor, is necessary for the development of gonadotrophs [24]. However, no essential corticotroph specific factor has yet been clearly identified. NeuroD, also known as Beta2, is a cell-restricted basic helixloop-helix (bHLH) transcription factor implicated in late neuronal differentiation [25] which has also been isolated as a cell-specific transcription factor of the insulin gene [26]. NeuroD is expressed in a number of tissues including pancreatic endocrine cells, the intestine, and brain including the mouse corticotroph AtT-20 cell line, but not the rat somatotroph pituitary cell line (GH3) [26]. Drouin and colleagues have recently demonstrated that NeuroD expression does appear to be restricted to corticotrophs in the mouse pituitary, and that NeuroD, in association with ubiquitous bHLH dimerization partners, specifically recognizes and activates transcription from the POMC promoter E box that confers transcriptional specificity of POMC to corticotrophs [27,28]. However, the necessity of this transcription factor for corticotroph specification or differentiation is still undetermined. Recently, a corticotroph-restricted transcription factor. TPIT, has been identified, which interacts with PTX1 and is required for POMC transcription, as discussed subse-

Chapter 2

quently. However, its role in corticotroph development is not known, and awaits further study. Because no instrinsic factor has yet been identified that is absolutely required for the development of corticotrophs in situ, it is possible that they arise elsewhere during fetal life and migrate into the anterior pituitary. One possible source could be neural crest, as this is the origin of POMC-producing melanocytes which migrate to the skin [29]. Extrinsic Signals

The oral ectoderm, from which Rathke’s pouch forms, and the neural ectoderm of the ventral diencephalon, from which the hypothalamus arises, are in direct contact at the time of the formation of Rathke’s pouch [30]. Classical explant experiments have indicated that inductive signals arising from mesenchyme and neural ectoderm adjacent to Rathke’s pouch are required for pituitary organogenesis and cell line specification [31]. The necessity for extrinsic signals for development of corticotrophs is supported by the demonstration that the expression of POMC in ectoderm explants requires coculture with mesoderm [32]. Direct evidence confirming the necessity of the ventral diencephalon for proper pituitary organogenesis has recently been obtained from examination of mice bearing null mutations in the Nkx2.1 homeobox gene, also known as T/EBP or TTF1 [33]. This gene is not expressed in the oral ectoderm or in the developing pituitary at any time during embryogenesis [34], but Nkx2.1-null mice demonstrate severe defects in the development of the diencephalon and also fail to develop a pituitary gland [33]. The nature of the extrinsic signals that promote pituitary organogenesis and cause cell-line specification is an area of active investigation. A number of secreted factors have been implicated in pituitary development and cell-line specification including bone morphogenic proteins (BMP4, BMP2), fibroblast growth factor 8 (FGF8), Sonic hedgehog (Shh) and Wnt5a [35–37]. A model of coordinate control of anterior pituitary progenitor cell identity, proliferation and differentiation imposed by FGF8 secreted from the dorsally located infundibulum and BMP2 from ventrally located mesenchyme has been proposed [37]. It has been suggested that corticotroph progenitors progress to a definitive corticotroph state only after escaping both FGF8 and BMP2 signaling [37]. While the precise signals and interactions required for pituitary organogenesis and corticotroph specification are incompletely understood, it is clear that signals from the mature hypothalamus like corticotropin releasing hormone (CRH) and vasopressin (AVP) are not required for corticotroph specification. The anterior pituitary appears to develop normally in a Crh-null mouse [38]. Furthermore, deletion of the class III POU transcription Brn-2 results in failed maturation of Avp-, Crh-, and oxytocin producing neurons of the hypothalamus and failed maturation of the posterior pituitary with no apparent defect in the maturation of any anterior pituitary cell type [39].

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Localization of Non-ACTH Peptides with Corticotrophs Many other neuropeptides have been found to be colocalized with ACTH within corticotrophs, although in most cases it is not clear whether this is due to synthesis or binding of the peptide within the cell. Neurophysin (but not vasopressin) colocalizes with ACTH in both normal and adenomatous corticotrophs [40]. Neurophysin immunocytochemical staining is especially prominent in corticotrophs in the zona intermedia which appear to project into the posterior pituitary. Chromogranin A has been described in the majority of corticotroph adenomas, although only a fraction of these patients have elevations of circulating plasma chromogranin A levels [41]. Galanin is present in all normal corticotrophs as well as in the majority of corticotroph adenomas which have been examined [42]. Galanin has also been found in those corticotrophs in the zona intermedia which appear to be migrating into the posterior pituitary. It has also been described in corticotrophs which have undergone Crooke’s hyalinization. Calpastatin, an inhibitor of the calcium-dependent cysteine proteases, calpain I, and calpain II, has been found in all ACTH-containing cells of the anterior pituitary, including those in the median wedge as well as those extending into the posterior pituitary [43]. Vasoactive intestinal peptide has been found in corticotroph adenomas, but only rarely in normal corticotrophs [44]. Normal corticotrophs contain small amounts of cytokeratin, whereas corticotrophs which have undergone Crooke’s hyalinization are markedly positive for this protein [45]. Corticotrophs appear to have few structural characteristics associated with neuronal cell types, for they are negative for neurofilament, vimentin, glial fibrillary acidic protein and desmon [45,46]. Likewise, corticotrophs in the zona intermedia, unlike those in the pars intermedia of rodents, appear not to be innervated by neurons, since they do not stain with any of these neuron-specific markers [47]. Similarly, synaptophysin, a 38 kD integral membrane glycoprotein found in presynaptic vesicles of neurons stains only weakly in corticotrophs [48]. These findings, together with a lack in humans of an anatomically discrete intermediate lobe with large numbers of a-MSH-containing cells, has led most investigators to conclude that there is no functional counterpart of the rodent pituitary intermediate lobe in humans.

Extra-pituitary Localization of ACTH and Related Peptides Although the vast majority of ACTH is synthesized in anterior pituitary corticotrophs, it is also expressed in several nonpituitary human tissues, both within and outside of the central nervous system. Within the central nervous system, ACTH and its related peptides are expressed to the greatest degree in cell bodies of the infundibular nucleus of the basal hypothalamus (analogous to the rodent arcuate nucleus) [49–51]. (See POMC Biosynthesis and Processing,

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pp. 52–53.) The cell bodies of these hypothalamic ACTH neurons are located at the base of the third ventricle, adjacent to the median eminence and pituitary stalk. These neurons project to limbic, diencephalic, mesencephalic, and amygdaloid sites [52]. Lesser amounts of ACTH and related peptides are found in substantia nigra, periventricular grey matter, and hippocampus [53]. Interestingly, POMC expression in the hypothalamus occurs in areas also known to express the orexigenic neuropeptides NPY and AGRP and project to many of the same hypothalamic targets [54,55]. The rodent arcuate nucleus prominently expresses the leptin receptor, implicating Pomc products in the regulation of appetite and energy homeostasis (see Melanocortin Receptor 4, p. 60) [56,57]. The brain areas containing ACTH also coincide with areas mediating stimulation analgesia, suggesting that expression of ACTH, or another product of the POMC gene (such as b-endorphin) in these sites may regulate pain perception [52]. Outside of the central nervous system, ACTH and other POMC gene products, including a-MSH and b-endorphin, are synthesized in a large number of human tissues, including in descending order of abundance, adrenal, testis, peripheral mononuclear cells, spleen, kidney, ovary, lung, thyroid, liver, colon, and duodenum [58,59]. In most of these tissues, POMC peptides are translated from truncated messenger RNAs lacking signal peptide sequences, suggesting that they cannot be secreted extracellularly [60,61], raising the question of their functional significance. However, adrenal and testis in addition express full length POMC mRNA, suggesting that these tissues may also secrete POMC-related peptides [58]. Recently, several additional cell types have been shown to produce POMC peptides including monocytes [62], astrocytes [63], gastrin-producing cells of the gastrointestinal tract [64], keratinocytes [65], skin melanocytes [66], and atrial myocytes [67]. GENE STRUCTURE ACTH is derived from a 266 amino acid precursor, proopiomelanocortin (POMC), so named because it encodes opioid, melanotropic, and corticotropic activities [68]. The human POMC gene is a single copy gene located on chromosome 2p23 [69]. It and the highly homologous opioid peptides, preproenkephalin A and preproenkephalin B (dynorphin), are all located on different chromosomes [70]. The human POMC gene is 8 kilobases (kb) long (Fig. 2.1). It consists of a promoter of at least 400 basepairs (bp) at the 5¢ end of the gene, followed by three exons, 86 (exon 1), 152 (exon 2), and 833 (exon 3) bp long, and two introns, 3708 (intron 1) and 2886 (intron 2) bp in length [69]. Exon 1 is not translated. Perhaps because of this, exon 1 of the human and other mammalian POMC genes are less than 50% identical. The initiator methionine is located 20 bp into exon 2, and is followed by a 26 amino acid hydrophobic signal peptide. Except for the signal peptide and 18 amino acids of the amino-terminal glycopeptide, the majority of

the POMC precursor is encoded by exon 3 [71]. Exon 2 is close to 90% identical between the POMC genes of humans and other mammals. Within exon 3 of POMC are located all known peptide products of the gene, including N-terminal glycopeptide, gamma-melanotropin (gammaMSH), joining peptide, a-MSH, ACTH, corticotropin-like intermediate lobe peptide (CLIP), b-lipotropin (b-LPH), b-MSH, and b-endorphin. Within exon 3, the regions encoding the N-terminal glycopeptide, a-MSH, ACTH, and b-endorphin, are greater than 95% identical between the human and other mammals [71]. In contrast, joining peptide, the region between the N-terminal glycopeptide and ACTH, is very poorly conserved among mammals [72], which has suggested to some workers that it does not encode a biologically important function [73].

POMC Gene Promoter Structure The promoter of the human POMC gene contains typical TATA and CAAT boxes 28 and 62 bp, respectively, upstream from the transcription start site. Using in vitro transcription [74] and transfection of the human POMC gene into heterologous cells [75], the POMC gene promoter has been shown to contain DNA elements which mediate increased transcription by cyclic AMP and decreased transcription by glucocorticoids. Although these DNA elements have not been precisely localized in the human POMC gene promoter, they are present with in the 700 bp 5¢ to the transcription start site [75]. The rat Pomc gene promoter has been studied more extensively. Drouin and coworkers, using DNA-mediated gene transfer into transgenic mice and tissue culture cells, found that the DNA sequences needed for corticotroph-specific expression and negative transcriptional regulation by glucocorticoid are contained within 543 bp of the transcription start site. These workers have reported the presence of a DNA sequence which mediates negative regulation by glucocorticoids (nGRE), located in the region of the CAAT box, which also binds nuclear proteins of the chicken ovalbumin upstream promoter (COUP) family. Using DNAase footprint and gel retardation analysis of the rat Pomc gene promoter, multiple synergistic DNA elements have been reported to be necessary for correct pituitaryspecific expression of the gene [76]. Recently, these workers have identified NeuroD, Ptx1, and Tpit as transcription factors which cooperate to cause corticotroph-specific expression of POMC [77].

POMC mRNA Transcription, Splicing, and Polyadenylation In addition to the major transcriptional initiation site present in corticotrophs, at least six other start sites have been found in several non-pituitary tissues (Fig. 2.1), including adrenal, thymus, and testes of the human [60] and rat [78]. These sites are all between 41 and 162 nucleotides downstream from the 5¢ and of exon 3. The mRNAs transcribed from

Chapter 2

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FIGURE 2.1. Structure of pro-opiomelanocortin (POMC) gene, messenger RNA (mRNA), and peptide products. (a) schematic diagram of gene (top), mRNA (middle) and POMC peptide (bottom). The gene consists of the promoter, 3 exons (thick bar, 1–3) separated by two exons (thin broken lines). Translated regions are stippled. The mRNA consists of 3 exons and the polyadenylate (AAAA) tail. The POMC precursor can be divided into 3 domains: I (N-terminal glycopeptide, joining peptide), II (adrenocorticotropic hormone; ACTH); and III (b-lipotropin; b-LPH). (b) Structure of individual products of POMC. Individual peptides are enclosed in different boxes depending on proteolytic cleavage specificities: = similar processing in pituitary and brain; = pituitary pattern of processing; = brain pattern of processing. = dibasic amino acids at proteolytic cleave sites. = glycosylation sites. = N-terminal acetylation sites. CLIP, corticotropin-like intermediate lobe peptide; a-MSH, a-melanocyte stimulating hormone.

these sites thus would be intronless, and the only truncated molecules that might be translated would be devoid of a signal peptide, and therefore could not be secreted [78]. Tissues containing these shorter forms of POMC mRNA, including adrenal, testis, spleen, kidney, ovary, lung, thyroid, and gastrointestinal tract, express ACTH, N-terminal glycopeptide1–61, and b-endorphin [58]. These truncated mRNAs are capable of being translated both in cell-free translation systems and in heterologous cells transfected with the appropriate fragment of the human POMC gene, although the peptide products are not secreted [61]. The significance of the expression of these truncated forms of POMC mRNA and their translated peptide products in human nonpituitary tissue is not clear.

A canonical polyadenylation signal is present in human POMC mRNA 23 bases upstream from the poly (A) addition site [79]. The length of the polyadenylate tail, attached to the 3¢ end of POMC mRNA and believed to play a role in translational efficiency of mRNA stability, is much longer in hypothalamic compared with pituitary cells in the rat [78], although this has not been studied in the human. GENE REGULATION In human anterior pituitary corticotrophs, POMC mRNA levels are increased by CRH and inhibited by glucocorticoids [80]. Similar results are seen in normal and adenoma-

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tous corticotrophs, although the glucocorticoid inhibition of POMC mRNA expression is less in the latter cells [80]. The same results are seen in rat anterior pituitary corticotrophs, where the opposing effects of CRH and glucocorticoids are due to their opposite influence on the transcription rate of the POMC gene [81]. Of interest, glucocorticoids have no effect on the POMC mRNA content of rat intermediate pituitary lobe corticotrophs [81]. CRH, acting via the type 1 CRH receptor [82] increases cAMP content in anterior pituitary corticotrophs [83]. CRH, vasopressin, and glucocotricoids all inhibit expression of CRH receptor 1 mRNA [84], which may limit the effect of these agents during the stress response, as discussed subsequently. The CRH-induced rise in cAMP is responsible for both the increase in POMC transcription and peptide synthesis as well as for the rise in intracellular calcium which results in ACTH secretion [85]. CRH mediates its stimulation of POMC transcription via the POMC CRH responsive element (PCRH-RE), which binds PCRH-RE binding protein [86]. The negative effect of glucocorticoids upon POMC gene transcription is thought to be mediated by a glucocorticoid–glucocorticoid receptor complex binding to cis-acting DNA sequences within the POMC promoter, although definitive evidence for this is lacking. The possibility exists that the glucocorticoid receptor complex does not bind directly to the POMC gene, but instead to another protein such as a positive transcription factor, and in this way mediates its negative effect on POMC gene expression [87]. Glucocorticoid stimulates, rather than inhibits, POMC gene expression [88] in the arcuate nucleus of the hypothalamus, the site of alphaMSH production. The significance of this is not clear, but could be involved in the inhibition of appetite by glucocorticoids in the postabsorptive state, when glucocorticoid levels are high. Recently, new insights into the development and regulation of the HPA axis have come from the discovery that leukemia-inhibitory factor (LIF) has an important role in these events [89]. LIF is a cytokine expressed in corticotrophs and folliculostellate cells beginning as early as 14 weeks gestation [90]. LIF stimulates transcription of POMC and expression of ACTH [90]. In many tissues, including pituitary, LIF expression is unregulated by inflammatory stimuli. The LIF receptor is a member of the class I cytokine receptor superfamily which heterodimerizes with gp130. In common with other family members, the LIF receptor signals through the Jak-STAT pathway, particularly utilizing Jak1 and STAT-3 [91]. As with other receptors coupled via these mediators, SOCS-3 inhibits POMC expression following its simulation by LIF [92]. Mice which overexpress a LIF transgene develop Cushing’s syndrome [93]. Their pituitary glands have corticotroph hyperplasia and multiple Rathke-like cysts lined by ciliated cells. Mice with targeted deletion of LIF have secondary adrenal insufficiency [94]. All of these data point towards LIF playing an important role in the regulation of ACTH secretion, perhaps most

importantly by immune and inflammatory stimuli [95]. In addition, the data suggest that LIF might be involved in the pathogenesis of Rathke-cleft tumors. Vasopressin potentiates the action of CRH on ACTH secretion, both in vitro [96] and in vivo [97]. Vasopressin’s effect is mediated by the vasopressin V1b (or V3) receptor and protein kinase C [98]. Despite this positive effect on ACTH secretion, vasopressin has been reported to decrease both basal and CRH-stimulated POMC mRNA levels in anterior pituitary cells [99]. b-adrenergic catecholamines, like CRH, also increase POMC mRNA levels in corticotrophs via a cAMP mechanism [100]. Insulin-induced hypoglycemia also causes an increase in POMC mRNA content in rat anterior pituitary corticotrophs [101,102], but whether this is secondary to an increase in hypothalamic CRH [101], vasopressin [103], or catecholamines [104] is not known. The inhibitory neurotransmitter GABA causes a decrease in POMC mRNA levels in intermediate, but not anterior pituitary corticotrophs [105]. POMC BIOSYNTHESIS AND PROCESSING The human POMC precursor has the potential to encode several overlapping peptides of biological importance (Fig. 2.1). Within the precursor, these peptides are separated from one another by two or more basic amino acids which serve as recognition sites for prohormone cleavage enzymes. In addition, POMC-derived peptides contain potential signals for amidation, glycosylation, acetylation and phosphorylation. Because the nomenclature of the various proteolytic products of POMC has been derived from both peptidemapping studies as well as molecular biological studies in which putative peptides had been predicted from inspection of nucleotide sequences [72], the terminology can be confusing. To avoid confusion, amino acid (aa) positions in this chapter are numbered as superscripts with reference to the 240 aa-long human POMC precursor, formed after removal of the 26 aa-long signal peptide. The 240 aa POMC precursor can be considered to be composed of three domains (Fig. 2.1). Domain I (aa 1–111), the N-terminal domain, encodes the 76 aa long Nterminal glycopeptide1–76 within its first 78 aa, and the 30 aa long C-terminal joining, or hinge, peptide ( JP79–108) within its last 33 aa. The middle Domain II (aa 112–152), 41 aa long, encodes the 39 aa ACTH112–150 peptide, which may be further processed to a-melanocyte stimulating hormone (MSH)112–124. The C-terminal Domain III, termed b-LPH153–240, is 88 aa long. It contains within it the 31 aa long b-endorphin210–240. Besides these peptides, several other products have been identified, although evidence for their existence and/or biological importance in man is not clear [106]. These include gamma1-MSH51–61, gamma2-MSH51–62, and gamma3-MSH51–76 in Domain I, corticotropin-like intermediate lobe peptide (CLIP)130–150 in Domain II, and gamma-LPH153–206 and b-MSH191–206 in Domain III (Fig. 2.1).

Chapter 2

Glycosylation of POMC Within the endoplasmic reticulum (ER), POMC undergoes initial glycosylation. Human POMC is glycosylated solely at two sites in the N-terminal glycopeptide1–76 of Domain I. Carbohydrate is added via an O-linked glycosylation to Thr45 and via an N-linked glycosylation to Asn65.

C-terminal Amidation of POMC Three human POMC products undergo C-terminal amidation [107]. These include N-terminal glycopeptide1–61, JP79–108, and -MSH112–124. In addition, these three products are also present in their Gly-extended forms, which may be incompletely-processed intermediates. C-terminal amidation is common among neuropeptides, and is usually essential for bioactivity [108]. This reaction is mediated by a bifunctional enzyme consisting of peptidylglycinea-amidating monooxygenese (PAM) and peptidly-ahydroxyglycine a-amidating lyase (PAL) activities, which transfer the amino group of a C-terminal Gly to the carboxyl group of the adjacent amino acid. Human PAM/ PAL exists in both a membrane-bound and free cytoplasmic form [109].

N-terminal Acetylation of POMC Two human POMC products undergo N-terminal acetylation. These are (as described below) a-MSH and bendorphin. In humans a-MSH exists predominantly in the nonacetylated form [110]. N-terminal acetylation of ACTH1–13-amide to form a-MSH results in increased melanotrophic activity [111] and decreased corticotrophic activity.

Proteolytic Processing of POMC POMC gives rise to several smaller, biologically active peptide products. These are generated by posttranslational cleavage of POMC by trypsin-like prohormone convertase endopeptidase enzymes which cleave the precursor on the C-terminal side of regions of two or more basic amino acid residues [112]. These basic amino acids are subsequently removed by a carboxypeptidase activity. Some POMC proteolytic products subsequently undergo amidation at their C-terminus or acetylation at their N-terminus, as described above. The posttranslational processing of POMC exhibits a remarkable degree of tissue-specificity, which recently has been postulated to be due to the differential distribution of processing enzymes in the various tissues which synthesize POMC (see below). Proteolytic Processing Enzymes

All proteolytic processing of human POMC occurs at either lys–arg or arg–arg residues (Fig. 2.1). Every lys–arg and arg– arg site within the human precursor is capable of being cleaved in vivo [106], whereas in the mouse and rat, additional arg–lys and lys–lys sequences at the N-termini of

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gamma1-MSH and b-MSH, respectively, appear to be utilized [106]. It is likely that the proteolytic digestion of human POMC at all sites is mediated by either of two structurally related endopeptidases, prohormone convertase 1 (PC1) [113] or prohormone convertase 2 (PC2) [113]. These enzymes, best studied in rodents thus far, are part of a seven member family of subtilisin/kexin-like mammalian proteinases [114], and are distributed specifically within endocrine cells and neurons. Both enzymes are capable of cleaving neuropeptide precursors, including POMC, proinsulin, and proglucagon, at dibasic sites, and each appears to manifest distinct preferences for different sites within the same precursor prohormone, with PC2 able to cleave at a wider selection of available dibasic sites than PC1 [115]. The tissue distributions of PC1 and PC2 mRNAs are distinctly different. PC1 is abundant in approximately 20% of anterior pituitary cells (presumably including corticotrophs), in all intermediate lobe pituitary cells (of the rodent), and in the supraoptic nucleus of the hypothalamus [116]. In contrast, PC2 is absent from the anterior pituitary, but is highly expressed in rodent pituitary intermediate lobe, multiple sites within the central nervous system, including cerebral cortex, hippocampus, and thalamus, and in pancreatic islet cells [116]. The differential tissue-specific distribution of these enzymes matches nicely with the known tissue-specific differences in POMC proteolytic processing (see below), suggesting that PC1 is responsible for the POMC cleavage products found in anterior pituitary corticotrophs, whereas PC2 cleaves POMC in pituitary intermediate lobe (of lower mammals) and in the brain. This suggestion is supported by studies of the differential processing of POMC by PC1 and PC2 [115,116]. Recently, a patient with severe childhood obesity and hyperproinsulinemia with postprandial hypoglycemia has been identified with compound heterozygous mutations in the PC1 gene [117]. The patient presented with multiple endocrine abnormalities including impaired glucose tolerance and postprandial hypoglycemia which has been attributed to the secretion of large amounts of proinsulin given its partial insulin-like action and long biological half-life. She also suffered from hypogonadotropic hypogonadism with primary amenorrhea, but otherwise normal development of secondary sexual characteristics. Ovulation was induced with exogenous gonadotropins, and the patient delivered healthy quadruplets. The pregnancy was complicated by gestational diabetes requiring insulin treatment. She also suffered from mild adrenal cortical insufficiency with complaints of fatigue and excessive daytime somnolence reversed by glucocorticoid administration. The adrenal cortical insufficiency was attributed to defective POMC processing with elevated levels of serum ACTH precursors confirming a role for PC1 in human POMC processing [117]. Following the generation of peptide products by prohormone convertase enzymes, the C-terminal basic amino acids are removed by carboxypeptidase activities. Carboxypeptidase E (Cpe) is required for the excision of paired

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dibasic residues of various peptide prohormone intermediates, including those derived from proinsulin and POMC [118]. The mutation in a strain of hyperproinsulinemic, late-onset obese mouse, the fat/fat mouse, has recently been mapped to the carboxypeptidase E gene [118]. A missense mutation has been identified in the Cpefat allele and these mice demonstrated a 20-fold decrease in Cpe enzymatic activity in pituitaries and isolated islets. The sorting of POMC into secretory granules is probably mediated by a signal “patch” [119], located within the tertiary structure of the molecule, which directs it to the granules. The presence of all POMC processed products in equimolar amounts within secretory granules suggests that this sorting precedes proteolytic cleavage of POMC. This suggestion is supported by data which demonstrates that initiation of proteolytic processing of POMC begins in the trans-Golgi system and continues in secretory vesicles, and is also consistent with data demonstrating localization of PC1 and PC2 within the TGN and/or dense core secretory granules [120]. Since POMC as well as PC1 and PC2 all contain signal peptides and are presumably colocalized throughout the endoplasmic reticulum, prohormone convertase activity must be inhibited in the endoplasmic reticulum and cis-Golgi regions. Proconvertase activity may be regulated by the local intracellular environment of these compartments and/or by control of catalytic activation of prohormone convertase. 7B2 is a small, hydrophobic acidic protein originally isolated from porcine and human anterior pituitary glands [121] that is widely distributed in neuroendocrine tissues [122] and found to associate specifically with PC2 [123]. The 27-kDa 7B2 precursor protein is cleaved to a 21-kDa protein and a small carboxy-terminal peptide (CT peptide) [124]. Interaction of 7B2, particularly the 21-kDa fragment, with proPC2 appears necessary for the generation of mature and active PC2 in the trans-Golgi region [125]. The 7B2 CT peptide is a nanomolar inhibitor of PC2 in vitro, but its role in vivo has not been defined. The role of 7B2 in activating proPC2 has been confirmed in vivo in 7B2-null mice [126]. 7B2-null mice are devoid of PC2 activity and deficiently process islet hormones and Pomc. The mice are hypoglycemic and demonstrate hyperproinsulinemia and hypoglucagonemia. These mice also demonstrate profound intermediate lobe ACTH hypersecretion with minimal conversion of this peptide to a-MSH, resulting in a severe Cushing’s syndrome that causes death by 9 weeks of age [126]. Curiously, PC2 null mice demonstrate similar islet cell dysfunction resulting in hypoglycemia, but do not abnormally produce a Cushingnoid syndrome [127]. This discrepancy suggests additional functional roles for 7B2, which are further suggested by the localization of 7B2 in regions of the brain lacking PC2 [128]. Recently, the suggestion that 7B2 may represent one member of a family of related convertase inhibitor proteins has been proposed with the identification of the protein proSAAS [129]. ProSAAS is a 26 kDa granin-like neuroen-

docrine peptide precursor isolated from rodents and humans with structural similarity to 7B2, including a proline-rich sequence in the first half of the molecule and a C-terminal peptide (SAAS CT peptide) following a dibasic cleavage sequence [129]. Overexpression of proSAAS in AtT-20 cells reduces the rate of Pomc processing and the SAAS CT peptide is a nanomolar competitive inhibitor of PC1, but not PC2 [129]. Tissue Specificity of POMC Processing

In human corticotrophs, POMC is processed predominantly into N-terminal glycopeptide1–76, joining peptide ( JP)79–108, ACTH112–150, and b-LPH153–240 (Fig. 2.1) [130]. Much smaller amounts of a-MSH112–124, CLIP130–150, bendorphin210–240, and a truncated form of N-terminal glycopeptide1–61, also known as “big” gamma-MSH, are also present [107]. There is no evidence for cleavage after arg50, and therefore no evidence for the presence of gamma1MSH51–61 in the human pituitary. JP79–108 exists as both a monomer and homodimer, most likely joined via disulfide bonding between the single Cys87 of two molecules [73]. Although JP79–108 has been postulated to stimulate adrenal androgen steroidogenesis [131], some studies do not support this hypothesis [73]. A function for N-terminal glycopeptide has also not been assigned, although one intriguing study reported that it is capable of stimulating aldosterone release from adrenal cells [130]. Whereas the production of distinct POMC peptide derivatives is clearly segregated between the anterior and intermediate lobes of the rodent, in the human, small, acetylated POMC peptide derivatives colocalize with larger desacetylated POMC peptides in corticotrophs of the anterior pituitary, suggesting that the strict dichotomy between corticotrope and melanotrope POMC processing observed in rodents and other species does not extend to human pituitaries [132]. Levels of desacetyl-a-MSH are elevated in pituitary corticotrophs [133] and plasma [134] of patients with Addison’s disease, Cushing’s disease, and Nelson’s syndrome. Desacetyla-MSH has approximately 75% of the melanotrophic activity as does a-MSH [111], whereas ACTH is only 5% as potent as a-MSH in this regard [111,135]. Because the serum levels of ACTH are 50- to 100-fold higher than levels of desacetyl-a-MSH in patients with Cushing’s disease, Addison’s disease, and Nelson’s syndrome, it is likely that the hyperpigmentation associated with these disorders is largely due to the melanotrophic effect of ACTH, and not MSH. Thus, in human anterior pituitary corticotrophs, the POMC precursor is predominantly cleaved at limited lys–arg sites into two peptides in Domain I (N-terminal glycopeptide1–76 and JP79–108), one peptide in Domain II (ACTH112–150) and one peptide in Domain III (b-LPH153–240) (see Fig. 2.1). As discussed above, human POMC is also expressed in several brain sites outside of the anterior pituitary, predominantly in the arcuate nucleus of the anterior hypothalamus. In these extra-pituitary locations, POMC is

Chapter 2

processed to a greater extent than in anterior pituitary. In brain, ACTH112–150 is cleaved to a-MSH112–124 and CLIP130–150 such that the amount of a-MSH relative to ACTH is 300-fold higher in hypothalamus, telencephalon, and mesencephalon than it is in anterior pituitary [136]. As in anterior pituitary, a-MSH is almost exclusively present in the desacetyl form [50]. Adding additional levels of complexity to the issue of tissue specificity of POMC processing are results from a dopamine D2 receptor (D2R) deficient mouse. Similar to dopaminergic tonic inhibitory control of prolactin expression in lactotrophs, POMC expression in the rodent intermediate lobe is under inhibitory dopaminergic control mediated via D2 receptors [137]. D2R-deficient mice demonstrate mild intermediate lobe hyperplasia accompanied by upregulation of both PC1 and PC2 [138]. These mice present with unexpectedly high levels of ACTH with corresponding adrenal hypertrophy and increases in corticosteroid secretion [138]. The altered prohormone convertase levels in these mice suggest the possibility of dynamic regulation of prohormone processing within specific tissues. HORMONE MEASUREMENT ACTH was one of the first substances to be measured by radioimmunoassay (RIA) [139]. In pioneering work using polyclonal antisera, Berson and Yalow described the measurement of ACTH and related peptides in normal persons and those with ectopic ACTH production by lung cancer [140]. These studies provided among the first data that ACTH was synthesized from larger precursors, which were termed “big” and “big–big” ACTH. The ACTH RIA can be performed on unextracted plasma, but suffers from a limit of sensitivity of approximately 25 pg/ml, and is therefore often unable to detect levels of plasma ACTH in the normal basal range. Radioimmunoassay remained the standard method for the measurement of plasma ACTH until the development of the immunoradiometric assay (IRMA). ACTH IRMAs employ two antibodies, one or both monoclonal, against ACTH. The solution-phase antibody is radiolabeled, and the solid-phase antibody is linked to a bead or other solid support. In general, the ACTH IRMA on unextracted plasma compares very favorably with RIA [141], being more sensitive, more reproducible, and more rapid [142]. Most IRMAs have lower limits of detection of 3–5 pg/ml and coefficients of variation of less than 10% up to 5000 pg/ml [141,142]. Depending on the antigenic specificity of the chosen antibodies, the ACTH IRMA may detect only intact ACTH [141], both ACTH and POMC precursor peptides [143,144], or only POMC precursors [145]. It is essential to know the sequence specificity of any IRMA in clinical use, for some, unlike most polyclonal RIAs, are incapable of detecting ACTH precursors which may be secreted by lung carcinomas [143]. Despite the wide availability of the ACTH IRMA, results from different laboratories are diffi-

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cult to compare because of lack of agreement on a suitable ACTH reference standard. An immunofluorometric assay (IFMA) for ACTH has also been developed [146]. Its sensitivity and accuracy are similar to those of the IRMA. Its principal advantage is its speed and the use of a nonradioactive label which is stable for over 1 year. Plasma assay of POMC products other than ACTH, including b-endorphin [147] and N-terminal glycopeptide, has been suggested as an adjunct in the evaluation of the hypothalamic-pituitary-adrenal (HPA) axis. In general, the levels of these hormones parallel that of ACTH in various HPA axis abnormalities [148]. However, except for use as a screening test for lung carcinoma associated with preferential secretion of N-terminal glycopeptide1–61 or proACTH1–150 [148], such tests are much less helpful than the ACTH IRMA because of the low concentration of other POMC peptide fragments compared with ACTH in most physiologic and pathologic settings. SECRETION OF ACTH: BIOCHEMISTRY AND PHARMACOLOGY Secretion of ACTH from the corticotrophs of the anterior pituitary is mediated by several factors (Fig. 2.2). CRH and vasopressin are the primary secretagogues for ACTH, although a number of other agents may also affect its release, and glucocorticoids are the major negative regulators of ACTH secretion. Once a ligand has bound to its receptor, release of ACTH from the corticortroph is mediated by second messengers through one of four signal transduction pathways, involving either protein kinase A, protein kinase C, glucocorticoids, or the Janus kinase/STAT system. (This

–

CRH

AVP

Catecholamines IL1β IL-6 TNFα

+

–

Cortisol

ACTH

Adrenal

+

+

LIF IL-6 IL1β TNFα

CRH IL1β IL-6 TNFα

FIGURE 2.2. Control of adrenocorticotropic hormone synthesis and secretion.

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last pathway, regulated by LIF, was described in a previous section, p. 50.) These pathways result in changes in the phosphorylation pattern of specific cellular proteins, and/ or in intracellular calcium levels, impacting on ACTH synthesis and release. Circulating ACTH then binds to receptors, primarily in the adrenal gland, leading to steroid biosynthesis.

Second Messenger Regulation of ACTH Secretion Protein Kinase A Pathway

CRH stimulates ACTH release via the cAMP-protein kinase A pathway. CRH stimulates adenylate cyclase activity, which then increases cAMP [83]. Forskolin, a direct stimulator of adenylate cyclase activity, and 8-bromo-cAMP, a cAMP analog, both markedly stimulate ACTH release, and increase CRH-stimulated ACTH release, but do not potentiate ACTH release from cells maximally stimulated by CRH [149]. The CRH-stimulated increase in cAMP activates cAMP-dependent protein kinase A [83]. Protein Kinase C Pathway

AVP V1b and V2 receptors are coupled to different second messenger systems. V1 receptors in the pituitary mobilize calcium through activation of phospholipase C, whereas V2 receptors activate adenylate cyclase, increasing cAMP [150]. AVP-stimulated ACTH release is mediated via the protein kinase C pathway [151]. Evidence that AVP acts through the protein kinase C pathway in the anterior pituitary includes the finding that AVP stimulates accumulation of inositol phosphates in rat anterior pituitary cells [152]. Phorbol esters, which activate protein kinase C directly by substituting for diacylglycerol and binding to protein kinase, induce ACTH release in rat pituitary cell cultures. Activation of protein kinase C appears more important in the sustained phase of AVP-stimulated ACTH release, not in the initial phase [153].

CRH CRH Stimulation of ACTH Secretion

Perfused human fetal pituitaries and cultured human fetal pituitary cells in culture secrete ACTH in response to CRH [2]. Rats have been used extensively in the study of ACTH secretion, as it is difficult to culture human corticotroph cells in vitro. In dispersed rat anterior pituitary cells, CRH stimulates a ninefold increase in ACTH release that is sustained for as long as the cells are exposed to CRH [83]. Even with maximal CRH stimulation, only 6% of cultured pituitary cells release ACTH [154]. CRH stimulates ACTH synthesis as well as release. In humans, the biphasic response may reflect secretion of a ready pool of ACTH, followed by later release of newly synthesized protein [155]. Exposing cells to CRH for a long period of time results in an increase in ACTH in the cell and in the medium. Thus, the sustained phase of ACTH

secretion may represent later release of newly synthesized ACTH peptide. Most of the CRH released into the hypophyseal blood is derived from the PVN [156]. Concentrations of oCRH that are similar to the concentrations of CRH found in rat portal plasma have been shown to increase secretion of ACTH in human fetal hemipituitaries in vitro [157]. The CRH content in the hypophyseal portal blood of anesthetized male rats is about 100 pM [158], which exceeds the in vitro threshold of 10–20 pM CRH to stimulate ACTH release [159]. In addition to stimulating ACTH expression and release, CRH can also directly stimulate secretion of glucocorticoids from the adrenal gland [160]. CRH Receptors

In human pituitaries, CRH has been shown to bind to sites in the anterior lobe [161]. The distribution of binding sites in human pituitary correlates with the distribution of corticotrophs [161]. CRH receptors in the anterior pituitary gland are low capacity, high affinity receptors, with a Kd for CRH binding of about 1 nM [162]. To date, two CRH receptor genes have been identified in humans and other mammals [163–165], with a third additional one being described in the catfish [166]. The type 1 receptor [163,165] is expressed predominantly in anterior pituitary corticotroph cells, whereas the type 2 receptor [164] is more widely distributed in the brain and periphery, particularly in cardiovascular tissue. The type 1 receptor binds and is activated by both CRH as well as the CRH-like peptide, urocortin [167]. This receptor mediates the actions of CRH at the corticoctroph. In addition, the type 1 receptor mediates fear and anxiety behaviors following stressors, even in CRHdeficient mice [168]. Mice with deletion of the CRH type 1 receptor gene show reduced fear and anxiety [169]. These data suggest that a CRH-related peptide, possibly urocortin or another unknown member of the CRH family, mediates fear responses via the CRH type 1 receptor. The CRH type 2 receptor binds urocortin with over 20-fold higher affinity compared to CRH. Its distribution, along with the hypotensive cardiovascular response to infused urocortin which is abolished in CRH type 2 receptor-deficient mice [170], suggests that the receptor may be involved in blood pressure control. This may underlie the hypotension observed during the CRH stimulation test. Recently, a nonpeptide, type 1-specific CRH antagonist has been developed which is orally active [171,172]. It should help to elucidate the role of CRH in ACTH regulation and other pathways.

AVP AVP Stimulation of ACTH Secretion

In rat anterior pituitary cells, AVP causes a twofold increase in ACTH release [173]. AVP elicits an initial rapid release of ACTH, observable within 5 seconds, reaching a maximum in less than 1 to 2 minutes, and lasting less than 3 to 6 minutes. This is followed by a second phase lasting

Chapter 2

for several hours. In humans, AVP infusion by itself has only a small effect on ACTH release. AVP is synthesized in the same parvocellular hypothalamic paraventricular nuclear neurons which express CRH, and appears to be coreleased with CRH at the median eminence into the portal hypophyseal system [174]. However, a substantial amount of AVP in the portal blood is released from projections from the supraoptic nuclei in the median eminence [156]. AVP from the posterior pituitary may also reach the anterior pituitary through portal vessels that connect the two [175]. This raises the possibility that increased vasopressin secretion from the posterior pituitary, in response to hyperosmolality, may also stimulate ACTH secretion. AVP Receptors

A single population of specific AVP receptors have been identified in rat anterior pituitaries, which are distinct from CRH receptors [176]. Most corticotrophs have AVP receptors, since 80% of the ACTH-secreting cells in the pituitary bind AVP [176]. The anterior pituitary AVP receptors are distinct from the V2 renal receptors and the V1 hepatic/pressor receptors [150]. This has led to the classification of hepatic/pressor receptors as V1a receptors and anterior pituitary receptors as V1b [150], or V3, receptors. V1a, V1b, and V2 receptors can be distinguished by their patterns of recognition of AVP analogues [177]. V1a binding sites in the rat anterior pituitary have a Kd of about 1 nM, and the minimal effective dose of AVP is 0.1 nM [173]. dDAVP (desmopressin), an AVP analog with V2 receptor affinity, has an insignificant effect on plasma ACTH levels, though it does increase, but is not additive to, CRHstimulated ACTH release [178]. During the past 6 years, the genes for all three vasopressin receptors (V1a, V2, and V1b) have been identified [179]. They are highly related members of the 7 transmembrane, G protein coupled, receptor family. As anticipated, V1b receptor mRNA is highly expressed in anterior pituitary corticotrophs, and is coupled to stimulation of POMC gene expression and ACTH secretion via the protein kinase C pathway [180,181].

Modulators of CRH and AVP Release CRH is the most important physiologic ACTH secretagogue [182]. Stressors, endogenous circadiam rhythms, and glucocorticoids influence CRH release [183]. In the rat, afferent inputs to the PVN may mediate the action of stressors by controlling the release of CRH [183]. Sources of neuronal afferents to the hypothalamus include the amygdala and hippocampus of the limbic system, and brainstem regions involved in autonomic functions [184]. In rats, acetylcholine, norepinephrine, angiotensin II, and possibly CRH itself, increase CRH concentrations in the hypophyseal portal plasma. On the other hand, AVP, b-endorphin, and GABA inhibit the ACTH response to stress [182,185–187].

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In rodents, central catecholamines stimulate ACTH release, and the effects of catecholamines on ACTH secretion appear to be mediated via secretion of CRH, and possibly AVP, into the hypophyseal portal circulation [188]. However, in humans, catecholamines have little direct effect on ACTH secretion from the pituitary. Peripheral catecholamines, increased by a variety of stresses, do not cross the blood–brain barrier to reach the hypothalamus, but do reach the pituitary, yet do not increase basal or CRHstimulated plasma ACTH levels. This suggests that the increased peripheral levels of epinephrine and norepinephrine generated during stress are probably not responsible for the increase in ACTH, and that catecholamines do not act directly on the pituitary to stimulate ACTH release [189]. In humans and other intact mammals, the impact of angiotensin II on basal or CRH-stimulated ACTH release is unclear. In humans, infusion of angiotensin II alone does not increase ACTH release [178]. However, in dispersed rat anterior pituitary cells, angiotensin II does stimulate release of ACTH [190]. Angiotensin II has a synergistic effect with CRH in stimulating ACTH release in humans in vivo [178]. Angiotensin II potentiates CRH-stimulate ACTH release from cultured anterior pituitary cells [191], although it is less effective than AVP, and potentiates the CRH-stimulated increase in cAMP. Angiotensin II and AVP do not potentiate each other’s effect on ACTH release, suggesting that they act via the same mechanism. Interleukin-1 beta stimulates ACTH release in conscious rats by acting on the hypothalamus [192] to induce secretion of CRH [193,194]. Interleukin-1 does not cross the blood–brain barrier [195], but activates noradrenergic neurons in the brainstem and hypothalamus, which may stimulate CRH secretion, especially in the median eminence. Prostaglandins may be involved in the response to interleukin-1, since ibuprofen, which blocks the formation of prostaglandins, blocks endotoxin-induced ACTH release in humans [196]. Tumor necrosis factor is a potent secretagogue for ACTH, and when administered to human subjects, leads to an increase in ACTH, cortisol, and AVP [197], but inhibits CRH-, AVP-, and angiotensin II-stimulated ACTH secretion [198]. Tumor necrosis factor may stimulate ACTH secretion by stimulating CRH release from the hypothalamus [199]. However, there is evidence that the site of action of tumor necrosis factor is peripheral to the pituitary and hypothalamus [199]. Bacterial endotoxin, when administered to human subjects, increases ACTH, cortisol, and AVP release [196]. Tumor necrosis factor may mediate the hormonal responses to endotoxin, since tumor necrosis factor levels increase after endotoxin administration [200]. Endotoxin does not increase ACTH release from cultured rat adenohypophyseal cells [201]. In rats and humans, interleukin-6 leads to ACTH secretion via CRH-dependent [202] and CRH-independent pathways, most likely via a prostaglandin-dependent pathway [203]. GABA inhibits the secretion ACTH by inhibiting

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the release of CRH and possibly AVP [184]. GABAnergic inputs into the PVN of the hypothalamus have been characterized in animals, including inputs from the hippocampus [184]. Opiates and opiate agonists decrease ACTH secretion, and may tonically inhibit ACTH secretion [204–211], although some studies demonstrate no effect of naloxone, an opiate antagonist, on the basal levels of cortisol [212,213]. Opiates inhibit CRH- and AVP-stimulated ACTH release, and different opiate agonists differentially effect CRH- versus AVP-stimulated release [214]. In humans, morphine blunts CRH-stimulated ACTH release without decreasing AVP or catecholamine levels [214]. Modulation of CRH-stimulated ACTH secretion by opiates most likely occurs at a level above the corticotroph [214]. Met-enkephalin analogs inhibit ACTH secretion, and controversy exists as to whether inhibition occurs at the hypothalamus or pituitary. In humans CRH-induced ACTH release is almost completely abolished with pretreatment with a met-enkephalin analog [215].

Synergism Between CRH and AVP Alone, AVP is a less important physiologic ACTH secretagogue than CRH [182]. AVP potentiates CRH-stimulated release of ACTH from cultivated rat anterior pituitary cells by twofold. When cells are first exposed to CRH, reaching the sustained phase of CRH-stimulated ACTH release, and AVP is then added, the usual initial spike of AVP-stimulated ACTH secretion is superimposed on top of the CRHinduced plateau of secretion. Despite continued exposure to AVP, ACTH secretion decreases down to the level of the plateau phase of CRH-stimulated secretion [216]. In cells exposed to AVP before CRH, CRH does not potentiate AVP-stimulated ACTH secretion [216]. This, together with the presence of AVP and CRH in the same parvocellular neurons of the PVN, suggests that the two neuropeptides cooperate to regulate ACTH release. However, this remains to be proven in humans, as there are no examples of persons with defects in AVP synthesis or release who have impaired ACTH secretion.

Oxytocin In humans, low-dose oxytocin perfusion decreases plasma ACTH and cortisol levels [217], and infusion of oxytocin completely inhibits CRH-stimulated ACTH release [218]. Oxytocin acts via a similar mechanism as AVP. Oxytocin binds competitively to AVP receptors in the anterior pituitary [219], but is much weaker than AVP at stimulating ACTH release [220]. Like AVP, oxytocin stimulates ACTH secretion through the protein kinase C pathway.

Glucocorticoids Glucocorticoids are the primary negative regulators of ACTH secretion. Glucocorticoids act on corticotrophs to

inhibit the secretion of ACTH induced by AVP and CRH, the synergism between CRH and AVP, and substances that provoke production of inositol phosphates and cAMP. Glucocorticoids’ negative impact on ACTH regulation is also due to their inhibition of the principal stimulators of ACTH, CRH and AVP. In the rat, glucocorticoid receptors are widely distributed in the brain, including the PVN. The PVN is a site for glucocorticoid negative feedback, since dexamethasone decreases the amount of basal CRH in the hypothalami explant and in the PVN, and the CRH response to secretagogues like serotonin and to stress. Glucocorticoids inhibit CRH release and decrease intracellular CRH in the rat PVN [182], and dexamethasone has a local effect on the hypothalamus, decreasing CRH mRNA expression [221], and preventing the rise in intracellular of CRH and AVP usually seen after adrenalectomy [183]. Glucocorticoids increase, and adrenalectomy decreases, the amount of GABA in the hypothalamus and the hippocampus [222]. Glucocorticoids appear to feedback to increase the GABA activity of the hippocampus and hypothalamus, and thus inhibiting CRH release [222]. In the anterior pituitary, glucocorticoid inhibition of ACTH secretion in vitro is mediated via glucocorticoid receptors, and lack of glucocorticoid effect in the intermediate lobe of the pituitary is most likely because functional receptors are not present in these cells. The negative effect of glucocorticoids on CRH-, AVP-, angiotensin II-, and norepinephrinestimulated ACTH release [223] is biphasic, which may reflect an initial inhibition of ACTH release, followed by an inhibition of POMC biosynthesis. Glucocorticoids inhibit POMC secretion, gene transcription and mRNA levels in the anterior pituitary. EFFECTS OF POMC-DERIVED PEPTIDES IN SKIN AND ADRENAL The melanocortin receptors are a family of seven transmembrane-spanning, G-protein coupled receptors that are activated by melanocortin derivatives of POMC including a-MSH and ACTH. Activation of all five receptors results in adenylate cyclase activity and cAMP production. Cloning of the melanocyte MSH receptor (melanocortin 1 receptor–MC1R) [224,225] and the adrenal ACTH receptor (melanocortin 2 receptor–MC2R) [225] were quickly followed by the cloning of three additional family members. The five known melanocortin receptors show distinct tissue distributions throughout the nervous system and periphery and distinct selectivities for the various melanocortin peptides. Prior to the cloning of this receptor family, the actions of melanocortins were primarily known through the effects of MSH on pigmentation and the effects of ACTH on glucocorticoid secretion from the adrenals. However, many additional roles including cognitive and behavioral effects, effects on the immune system, and effects on the cardiovascular system have also been attributed to the melanocortins.

Chapter 2

With the cloning of this family of receptors, the physiologic roles of ACTH, MSH and other melanocortin derivatives are beginning to be elucidated.

Ligand Specificity The pharmacology of melanocortin receptor activation with a large number of natural and synthetic melanocortin peptides is the subject of extensive investigation. All five melanocortin receptors are activated by ACTH. However, MC2R binds only ACTH and is not activated by other melanocortin peptides [226]. The synthetic agonist 4norleucine, 7-D-phenylalanine-a-MSH (NDP-MSH) is the most potent agonist of MC1R, MC3R, MC4R, and MC5R [227,228]. The endogenous non-ACTH melanocortin peptides generally bind the melanocortin receptors with an order of potency MC1R > MC3R > MC4R > MC5R when expressed in COS cells and measurements are obtained in competition with NDP-MSH [228–232]. g-MSH is relatively selective for MC3R over MC4R and MC5R [228,233]. Whether the differences in melanocortin receptor specificity for different melanocortin ligands are physiologically relevant are unknown. Interestingly, Org 2766 and BIM 22015, two ACTH4-10 analogs, have no activity at any of the cloned MCRs, but have potent effects on central and peripheral nervous systems suggesting the possibility of undiscovered melanocortin receptors [234,235].

Melanocortin 1 Receptor The MC1R was initially cloned from primary melanoma tumors [224,236]. MC1R gene expression has also been confirmed in primary human melanocytes by northern blotting [236]. Other cutaneous cells including keratinocytes [237] and dermal fibroblasts [238] have also been reported to express MC1R, although the presence of MC1R mRNA [239] and MC1R protein [240] in keratinocytes could not be confirmed in other studies. Recent evidence demonstrates that the MC1R is expressed in keratinocytes and that its expression is induced by calcium and UV light treatments [241]. Corresponding to its expression in the skin, its function is perhaps most fully understood with respect to its role in cutaneous pigmentation (for review see [242]). Activation of melanocyte MC1R, via activation of adenylate cyclase, stimulates tyrosinase activity, the rate-limiting enzyme in melanogenesis [239]. The activation of tyrosinase results in an increased proportion of eumelanin (brown–black pigment) formation over pheomelanin (red–yellow pigment) formation resulting in increased pigmentation [243]. Mutations and variant alleles in the MC1R gene have been linked to variation in mammalian pigmentation. The extension locus has long been known to regulate pigmentation in mammalian species. The extension locus has been shown to encode the mouse MC1R [244]. The recessive yellow allele (e) at this locus results from a frameshift that produces a prematurely terminated, nonfunctioning recep-

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tor [244]. The sombre (Eso and Eso–3j) alleles and tobacco darkening (Etob) alleles, which have dominant melanizing effects, result from point mutations that produce constitutively active or hyperactive receptors [244]. The human MC1R is highly polymorphic, and mutant alleles have been associated with fair skin and blond or red hair [245–247]. Analysis of five naturally occurring common variants of MC1R associated with fair skin and red hair have revealed decreased stimulation of cAMP synthesis with no changes in or only slightly reduced aMSH binding [248]. In addition, MC1R variants have recently been shown to determine sun-sensitivity in humans with dark hair [249]. Melanocytes, which express POMC and several of its processed peptides [250,251], are derived from the neural crest [29]. It is likely that a-MSH produced via POMC expression in melanocytes is responsible for the pigmenting effects of ultraviolet light, and together with variations in MC1R discussed above, for the different degrees of skin pigmentation observed among ethnic groups. MC1R mRNA [252] and protein [253] have been detected in Leydig cells of the testis, luteal cells of the corpus luteum, and in the placenta. In the central nervous system, detection of MC1R mRNA and protein has been confined to a few scattered neurons of the periaqueductal grey in rat and human brains [254]. With the cloning of the melanocortin receptors, the role of melanocortins as anti-inflammatory agents has gained renewed interest [255–257]. Systemic administration of a-MSH has been shown to be beneficial in animal models of arthritis [258], adult respiratory distress syndrome (ARDS) [259], and septic shock [260]. Consistent with this role, MC1R expression has been documented on macrophages and monocytes [261,262]. Treatment of activated macrophages with a-MSH resulted in decreased nitric oxide production by inhibiting the induction of nitric oxide synthase II [262]. Lipopolysaccharides (LPS) and interferon treatments of neutrophils induced increases in neutrophil MC1R mRNA and treatment with a-MSH inhibited neutrophil chemotaxis by a cAMP dependent process [263]. Constitutive expression of MC1R has also been found on dermal microvascular endothelial cells where its expression can be increased by stimulation with IL-1b or a-MSH itself [264]. A potential role for melanocortins to regulate local inflammation in the brain has been proposed based on the evidence that tumor necrosis factor (TNF)-alpha secretion by an anaplastic astrocytoma cell line (A-172) is decreased by aMSH and that these cells express MC1R [63].

Melanocortin 2 Receptor—the ACTH Receptor Eary in the study of receptor biology, Haynes demonstrated the action of ACTH in generating cyclic adenosine monophosphate (cAMP) in adrenal cells [265]. The ACTH receptor was the first receptor that was shown to bind its ligand with high affinity and specificity [266]. In human adrenal glands, the ACTH receptor has a Kd of 1.6 nmol/l

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and about 3500 sites/cell [267]. The ED50 of ACTH for cAMP production is 0.11 nmol/l, 20-fold less than the Kd for binding [267]. The ED50 of ACTH for cortisol production is 2 pmol/l, 720-fold less than the Kd for ACTH binding, and 35-fold less than the concentration of ACTH needed to obtain a half-maximal increase in cAMP production [267]. Only a small percentage of ACTH receptors need to be occupied to achieve a maximal effect on steroidogenesis, which occurs at an ACTH concentration of 0.01 nmol/l [268,269]. Because the adrenal gland may express more than one of the five melanocortin receptors (see below) it is not possible to assign the biochemical characterizations to a specific melanocortin receptor. The human ACTH receptor (MC2R) was cloned based on its homology to MC1R [225]. In situ hybridization with the adrenal gland of the rhesus monkey demonstrated the presence of messenger RNA in the zona glomerulosa and fasciculata cells, and a weaker signal in the zona reticularis [225]. Initial attempts at characterizing the ligand specificity and cAMP signal generation in response to ACTH and other melanocortins was confounded by either poor levels of expression or the presence of endogenous melanocortin receptors in transfected cells [225,270,271]. Recently, the human MC2R has been stably transfected into the Y6 cell line, a mutant derived from the mouse Y1 adrenocortical cell line, that fails to express any endogenous MC2R [272]. Y6 cells alone demonstrated no cAMP response to micromolar ACTH, and MC2R transfected cells displayed an EC50 of 6.8 nmol/l [272].

Adrenal Actions of Acth The critical role of ACTH in regulating the synthesis and secretion of steroids from the adrenal gland has long been recognized. In adrenocortical cells, ACTH regulates lipoprotein uptake by receptor-mediated endocytosis from the plasma to lipid droplets by stimulation of lipoprotein receptors [273]. Within the lipid droplets, ACTH regulates hydrolysis of cholesterol esters by activation of cholesterol esterases or suppression of cholesterol acyltransferase, through cAMP-dependent protein kinase [274]. ACTH stimulates the transport of cholesterol to the mitochondria [275–277], principally via stimulation of steroidogenic acute regulatory protein (StAR) [278]. The rate-limiting step in steroidogenesis is the side-chain cleavage of cholesterol to pregnenolone, and is catalyzed by cytochrome P450-side chain cleavage enzyme [279] in the inner membrane of mitochondria of the adrenal [280], probably on the matrix side [281]. ACTH stimulation results in long-term and short-term effects on steroid hormone biosynthesis in the mitochondria [282]. The long-term effect of ACTH leads to an increase in the amounts of steroid hormone enzymes by increasing transcription of these genes [282]. StAR is the key protein which regulates cholesterol transport into the mitochondion [278]. Mutations in this protein result in defects in adrenal and gonadal steroidogenesis, which had been previously attributed to defects in side chain cleavage

enzyme activity [278]. Beginning several hours after ACTH administration, ACTH increases the levels of steroidogenic enzyme mRNAs in primary cultures of human adrenals by several-fold, including cholesterol side chain cleavage enzyme, 17-a-hydroxylase/17,20-lyase, 11-b-hydroxylase/ 18-hydroxylase/18-methyl-oxidase, and 21-hydroxylase cytochrome P-450 enzyme [283]. ACTH has a positive regulatory effect on its own receptors, and on the cAMP response to binding of ACTH to the receptor [284]. With sustained stimulation, down-regulation does occur, but physiologically this effect is minor since ACTH causes proliferation as well as steroid secretion. In addition to its prominent role in regulating adrenal steroidogenesis, ACTH exerts profound trophic effects upon the adrenal. Hypophysectomy results in adrenal atrophy and ACTH replacement restores adrenal gland weight in a dosedependent manner [285–288]. While the role of ACTH in adrenal hypertrophy is well established, its role in adrenocortical mitogenesis and hyperplasia is incompletely understood. The absence of ACTH induces apoptotic cell death in the adrenal cortex [289]. Prolonged ACTH administration not only blocks apoptosis, but also increases the adrenal DNA content in the rat [290], ACTH increases mRNA levels for c-fos and b-actin, proteins involved in cellular proliferation [291]. However, ACTH paradoxically inhibits mitosis of adrenocortical cells in culture [292–294]. ACTH antiserum administered to intact rats caused a highly significant decrease in corticosterone levels, but had no effect on adrenal weight [295]. Furthermore, ACTH inhibits the rapid compensatory proliferation of the remaining adrenal that normally occurs after unilateral adrenalectomy [296]. Another anterior pituitary-derived candidate for the stimulation of adrenal proliferation is the 28 amino acid Nterminal pro-opiomelanocortin peptide (N-POMC). This peptide is mitogenic in vitro and in vivo for the adrenal cortex, and N-POMC antiserum significantly diminishes adrenal mitotic activity after enucleation [297,298]. The compensatory adrenal growth that occurs after unilateral adrenalectomy may be mediated by a neural reflex that includes afferent neurons originating from the disrupted adrenal gland, the ventromedial nuclei of the hypothalamus, and efferent neurons innervating the remaining gland [299,300]. In 1947, Albright coined the term adrenarche to denote the developmental increase in adrenal androgens that occurs several years before the onset of gonadal maturation [301]. Adrenal androgen secretion may be sufficient for the development of some secondary sexual changes, including the development of pubic and axillary hair [302] and the maturation of sebaceous glands [303]. A condition in which axillary and pubic hair develop prematurely as a result of early adrenal androgen secretion has been termed premature adrenarche [304–309]. The mechanisms controlling adrenarche have remained obscure. Many hypotheses have been advanced, and many factors including ACTH, estrogens, prolactin, gonadotropins, growth hormone, glucocorticoids, androgens, and other POMC-derived products have been

Chapter 2

suggested as modulators of adrenal androgen secretion (for review see [310]). ACTH is widely accepted as a modulator of adrenal androgen secretion although, after administration of corticotropin, increases in dehydroepiandrosterone (DHEA) and dehydroepiandrosterone sulphate (DHEAS) tend to be small [311–313]. Furthermore, the increase in adrenal androgens that occurs in adrenarche is not accompanied by an increase in serum cortisol levels, leading to the suggestion that factors other than ACTH are responsible for adrenarche [307,309,314,315]. However, cortisol production rates do increase at the same time as does the increase in adrenal androgen secretion, suggesting a possible common link that could be due to ACTH stimulation of both steroid pathways [316]. Dissection of the physiology of adrenarche has proven difficult as the only animal showing an adrenarche similar to that of humans is the chimpanzee [317]. Recent data from patients with genetic defects in MC2R provide definitive evidence of the participation of this receptor in the process of adrenarche, as discussed subsequently. Familial Glucocorticoid Deficiency Syndrome

The familial glucocorticoid deficiency (FGD) syndrome is a rare autosomal recessive syndrome originally described by Shepard et al. in 1959 [318]. Patients typically present with symptoms resulting from their glucocorticoid insufficiency including hyperpigmentation, hypoglycemia, lethargy, and weakness [318–321]. The clinical course may be complicated by frequent infections [318,319]. Patients with FGD do not present with symptoms related to salt wasting, dehydration, or electrolyte disturbances, as the renin-aldosterone axis is preserved. The preservation of the renin-aldosterone axis clearly distinguishes this syndrome from childhood Addison’s disease. An unexplained feature of the syndrome is that many of the patients are reported to be unusually tall [318,319,321–325]. Patients with FGD typically have low or undetectable cortisol concentrations, but occasionally low, normal cortisol values, which respond subnormally to exogenous corticotropin [318,319,321–325]. Plasma ACTH levels are markedly elevated, often greater than 1000 pg/ml. The adrenal glands are atrophic, and only occasional cortical cells remained in the zona glomerulosa with no remnants of the zona fasciculata or reticularis, but the adrenal medulla appears normal [318,319,323]. This adrenal atrophy substantiates the physiologic relevance of ACTH’s trophic action on the adrenal gland. Recently, an examination of 11 patients with FGD revealed a discrepancy between partial glucocorticoid deficiency and significantly diminished DHEAS secretion confirming a significant contribution for ACTH (or at least for MC2R) in the onset of adrenarche [326]. A number of different homozygous or compound heterozygous missense and nonsense mutations in MC2R have been reported in patients with FGD, and in all cases, these mutations cosegregate with disease in the affected families [322,327–331]. Transient expression of the S741 mutant MC2R (ser74 Æ ile) in COS-7 cells resulted in a EC50 for cAMP production of 67 nmol/l, approximately 12-fold

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higher than the EC50 of the normal MC2R of 5.5 nmol/l. Expression of a Asp107 Æ Asn mutation in Cloudman S91 melanoma cells resulted in a six- to ninefold increase in the EC50 for cAMP generation. These results were confounded by endogenous melanocortin receptors making it difficult to determine whether the mutant receptor demonstrated defective ligand binding or defective coupling to adenylate cyclase [328]. Not all cases of the FGD syndrome can be accounted for by mutations in MC2R. Several families exhibiting the classic FGD signs and symptoms have no mutations in the coding regions of the MC2R gene [322,328,332,333]. These patients are clinically indistinguishable from patients with MC2R mutations except that they do not demonstrate increased height [334]. Recently, linkage of this form of FGD to a 12-cM region surrounding MC2R has been excluded in six families [332]. FGD, with ACTH resistance, also occurs in the Allgrove, also known as triple A (adrenal insufficiency, achalasia, and alacrima) syndrome [335]. Mutations in MC2R do not cause the resistance to ACTH. Recently, a novel gene, AAAS, has been identified on chromosome 12q13 in which several homozygous or compound heterozygous loss-of-function mutations are associated with this syndrome, in nine unrelated families [336,337]. This protein must in some way interfere with the expression or function of MC2R. ACTH-independent Activation of ACTH Receptor Pathways

Several rare causes of ACTH-independent Cushing’s syndrome are due to ACTH-independent pathological activation of ACTH receptor pathways. In the McCune-Albright syndrome, a mutation in the GTPase region of the stimulatory alpha subunit G protein, Gsa, can result in constitutive activation of protein kinase A in the adrenal cortex, leading to hypersecretion of cortisol and adrenal adenoma formation [338,339]. Ectopic expression of several other transmembrane G protein-coupled receptors, including those which bind gastric inhibitory polypeptide, luteinizing hormone, vasopressin, and catecholamines, in the adrenal cortex, may occur [340]. In this situation, Cushing’s syndrome results from the cognate hormone binding to the ectopically expressed receptor, leading to stimulation of adenylate cylase, activation of cyclic AMP, and cortisol hypersecretion [340]. Finally, patients with mutations in the regulatory subunit of protein kinase A, R1a, may develop Cushing’s syndrome secondary to micronodular adrenocortical hyperplasia [341,342]. This disorder, termed Carney complex, may be associated with myxomas of the cardiac atria and other tissues, and freckling of the skin [343]. EFFECTS OF POMC-DERIVED PEPTIDES OUTSIDE OF ADRENAL ACTH binds with high affinity to rat adipocytes and has potent lipolytic effects [344–347]. High levels of MC2R mRNA expression have been demonstrated in all murine

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adipose tissues examined, but MC5R mRNA expression was also found in a subset of these tissues [348]. Both MC2R and MC5R mRNA were identified in the 3T3-L1 cell line after these cells were induced to differentiate into adipocytes. The physiologic importance of the actions of melanocortins on adipose tissue is unclear. Primate adipose tissues have been reported to be insensitive to the lipolytic actions of ACTH [349]. MC2R mRNA expression was not detected in human adipose tissue [252]. In order to examine whether the MC2R might play roles in the hypothalamic–pituitary–adrenal axis in sites other than the adrenal gland, MC2R expression has been examined in the hypothalamus and the pituitary. MC2R mRNA could not be detected in the hypothalamus or the pituitary [350]. The expression of an ACTH receptor on human mononuclear leukocytes was suggested after a finding that high-affinity ACTH binding did not occur in a patient with FGD [351]. However, this finding is puzzling in retrospect as MC2R mRNA expression has not been demonstrated in leukocytes. Human skin cells express MC2R along with mRNA for three obligatory enzymes of steroid synthesis, the cytochromes P450scc (CYP11A1), P450c17 (CYP17), and P450c21 (CYP21A2) [352]. Slominski et al. have hypothesized that an equivalent of the HPA axis composed of locally produced CRH, CRH-receptor, POMC, and cortisol operate in mammalian skin as a local response to stress [353]. Recently, it was demonstrated that ACTH can induce DNA synthesis and cell proliferation in an oral keratinocyte cell line [354].

Melanocortin 3 Receptor Expression of the third cloned melanocortin receptor, MC3R, was originally identified in the brain, placenta, and gut [355]. Subsequently, MC3R expression was also documented in the heart [252]. Detailed mapping of the MC3R in the central nervous system by in situ hybridization localized the highest densities of MC3R mRNA to the hypothalamus and limbic system [356]. MC3R mRNA was found in the arcuate nucleus [356], the site of POMC expression within the hypothalamus. An autoradiographic approach using the non-selective [125I]NDP-MSH in competition with the relatively MC3R selective g-MSH or an MC4R selective synthetic agent HS014 has been used to visualize the MC3R and MC4R distributions within the central nervous system [357]. In the nucleus accumbens shell, the medial preoptic area, and the ventromedial nucleus of the hypothalamus, the MC3R dominates. In the lateral septum and the olfactory tubercle, both MC3R and MC4R seem to be present. The lack of overlap between the autoradiographic data and the data concerning MC3R mRNA expression may indicate the presence of MC3R on nerve terminals projecting from the arcuate nucleus. A physiologic role for MC3R has not yet been identified.

Recently, in a murine model of experimental gout, systemic treatment of mice with ACTH4–10 inhibited neutrophil accumulation [358]. This effect was blocked by he melanocortin receptor type 3/4 antagonist SHU9119. MC3R, but not MC4R, mRNA was detected in murine macrophages suggesting that MC3R may play a role in modulating inflammation. Recently, a mouse with targeted deletion of the MC3R gene has been created [359]. Although this mouse has a normal body weight, it has an increased fat to lean weight ratio, which seems to be due to an increased efficiency of converting ingested food into stored fuel. Thus, a function of melanocortins acting via MC3R may be to promote the conversion of energy in food into either lean body mass or forms of energy other than fat. This raises the possibility that in humans, mutations in MC3R may contribute to the “thrifty” genotype, as is found in Pima Indians [360].

Melanocortin 4 Receptor The MC4R is localized to the brain [361] and, in contrast to the MC3R, its expression has been documented throughout the central nervous system including the cortex, thalamus, hypothalamus, brainstem, and spinal cord [362]. In humans, the MC4R has not been detected in any peripheral tissue [252], although this distribution does not hold true for all animals as the MC4R is expressed in many peripheral tissues of the chicken [363]. The MC4R has garnered considerable attention as this melanocortin receptor subtype appears to play a central role in weight regulation. Mice homozygous for an Mc4r-null allele demonstrate autosomal dominant, maturity-onset obesity, hyperphagia, hyperinsulinemia, and hyperglycemia [364]. Heterozygotes have an intermediate phenotype. The Mc4r-null mouse also demonstrates increased linear growth [364], a feature unique to agouti yellow mice (discussed subsequently) and MC4Rnull mice among rodent obesity models. The MC4R has been implicated in leptin signaling as administration of the melanocortin receptor antagonist SHU9119 can block the reduction of food intake induced by central administration of leptin [365]. Leptin receptors are found on POMC expressing neurons in the arcuate nucleus and administration of exogenous leptin does not reverse obesity in Mc4rnull mice [366]. The role of MC4R in weight regulation in humans has been confirmed with the identification of individuals with dominantly inherited obesity segregating with mutations in the MC4R gene that result in frameshift errors [367–369]. Recent studies suggest that haploinsufficiency of MC4R may be a frequent, but incompletely penetrant, cause of human obesity [370,371]. The incomplete penetrance is highlighted by the absence of obesity in individuals with large deletions of chromosome 18q, a region that spans the MC4R gene [372]. A dominant-negative effect of mutant MC4R has been proposed, but cotransfection studies of mutant and wild-type MC4R in vitro showed that mutants

Chapter 2

affected neither signaling nor cell surface expression of wildtype MC4R [373].

Melanocortin 5 Receptor Expression of the fifth melanocortin receptor was originally recognized in the brain [374]. Subsequently, MC5R expression was documented widely at low levels in tissues including the adrenal glands, skin, adipocytes, skeletal muscle, kidneys, lung, stomach, liver, spleen, thymus, lymph nodes, mammary glands, ovary, pituitary, testis, and uterus [64,374–377]. In situ hybridization studies showed that within the adrenal, MC5R is predominantly expressed in the aldosterone-producing zona glomerulosa cells [376]. High levels of MC5R mRNA expression have been documented in the secretory epithelia of a number of exocrine and endocrine glands including Harderian, preputial, lacrimal, sebaceous, and prostate glands and pancreas [378,379]. Melanocortins have been reported to affect a number of exocrine glands. Removal of the neurointermediate lobe of the pituitary reduces sebaceous lipid production, and this reduction is restored by administration of a-MSH [380–382]. Exogenous ACTH and MSH increase secretion from the lacrimal gland [383,384]. Deletion of the murine Mc5r resulted in the loss of detectable binding of [125I]NDP-MSH to Harderian, lacrimal, and preputial glands, and skeletal muscle indicating that MC5R is the predominant melanocortin receptor in these tissues [378]. Development of the Mc5r-null mouse confirmed a physiologic role for the melanocortins in regulating exocrine gland function; the mice demonstrate severe deficits in water repulsion and thermoregulation as a result of decreased sebaceous lipid production [378]. MC5R also appears to be essential for hormonally regulated release of porphyrins from the Harderian gland [378]. The preputial gland is a specialized sebaceous gland implicated in pheromone production [385]. Exogenous a-MSH stimulates the release of a preputial odorant into the urine of male mice which stimulates aggressive attacks [386]. Chen and colleagues have hypothesized the existence of a hypothalamic– pituitary–exocrine axis which might provide a mechanism by which stress could alter behavior via the regulation of olfactory cues [378].

Diseases Caused by Melanocortin Receptor Ligands POMC Deficiency

Two individuals have been identified with genetic defects in the POMC gene [387] (Fig. 2.3). The first patient was a compound heterozygote for two mutations in exon 3 which interfere with appropriate synthesis of ACTH and a-MSH. The second patient was homozygous for a mutation in exon 2 which abolishes POMC mRNA translation. Both patients presented with early-onset obesity, adrenal insufficiency, and red hair pigmentation. The brother of patient one had died at the age of 7 months of hepatic failure following severe

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cholestasis and was found to have bilateral adrenal hypoplasia in the postmortem examination. All other anterior pituitary-derived hormones were normal. No symptoms related to b-endorphin deficiency have been noted. The heterozygous parents were normal in both families. Isolated ACTH deficiency of the pituitary due to other causes is also rare. It most commonly appears to be acquired in later life, and is more common in men [388–391]. Most cases have been reported in Japan, and are often associated with other autoimmune endocrinopathies, including Hashimoto’s thyroiditis and type 1 diabetes mellitus. It is likely that only pituitary ACTH, and not hypothalamic aMSH, is affected in these patients, as they are not obese and do not have red hair, unlike patients with POMC gene mutations [387]. Recently, the gene Tpit has been identified as a transcription factor specific to pituitary corticotrophs and melanotrophs, and absent in hypothalamic neurons which also express POMC [77]. It interacts with the POMC-specific transcription factor, Ptx1, to regulated POMC transcription. Two families with recessive mutations in TPIT have been described [77]. Consistent with its sites of expression, these patients have congenital, secondary adrenal insufficiency, but not obesity or red hair. Recently, a POMC-deficient mouse has been produced whose phenotype is similar to the human POMC-deficient syndrome and confirms the known functions of melanocortins [392]. The phenotype includes obesity, increased body length, yellow pigmentation, deficits in sebaceous gland function and thermoregulation, and adrenal hypoplasia and glucocorticoid deficiency. Adrenal glands could not be identified. In addition to undetectable corticosterone levels, aldosterone levels were also undetectable. The mutant mice lost 40% of their excess weight after 2 weeks of treatment with a stable a-MSH agonist. Although some of this weight loss was clearly attributable to decreased food intake, these same authors have clearly shown that a-MSH administration has additional lipolytic effects and results in increased energy expenditure [393]. Agouti and Agouti-related Protein

The agouti gene locus was identified 45 years ago as a genetic locus that controls the amount and distribution of eumelanin (brown–black) and pheomelanin (yellow–red) pigmentation in the mammalian coat [394]. However, analysis of mutations at the agouti gene locus have occupied investigators for nearly a century. The lethal yellow mutation at the agouti locus was the first murine embryonic lethal mutation and the first murine obesity syndrome to be characterized [395,396]. Agouti is a small 131 amino acid protein that is secreted by dermal papillae cells and acts to block melanocortin action on follicular melanocytes at Mc1r in the mouse [397,398]. The recombinant murine agouti protein is a potent nanomolar competitive antagonist for melanocortin receptors at Mc1r and Mc4r, relatively weaker at Mc3r, and only a micromolar inhibitor of Mc5r [397,399,400] and

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SECTION 1

Girl's weight

Boy's weight

35

–P97 82

Patient

–P90

30

72

25

–P75 –P50 –P25 –P10 –P3

62

20

–P97 –P90 –P75 –P50 –P25 –P10 –P3

15

52

Patient

42 32

10 22 5

0

12

0

3 6 9 12 18 24 30 36 42 48 Age (months)

2

0

2

4

6

8 10 12 14 16 18

Age (years)

represents the first known endogenous antagonist for a Gprotein coupled-receptor. In wild-type rodents, agouti expression is restricted to the hair follicle [398]. The human agouti gene which is 85% identical to the mouse gene is expressed much more widely including in adipose tissue, testis, ovary, and heart, and at lower levels in liver, kidney, and foreskin [401,402]. The human agouti protein, the agouti-signaling protein (ASIP), displays a similar pharmacologic profile for antagonism of melanocortin receptors as compared to the murine agouti protein except that ASIP may display both competitive and noncompetitive antagonism at MC4R and also appears to act as a noncompetitive antagonist of MC2R [403,404]. The heterozygous lethal yellow agouti mouse is characterized by a yellow coat-color, late onset obesity associated

FIGURE 2.3. Patients and weight curves of two unrelated patients with homozygous mutations in POMC. Note the obesity coupled with fair skin and red hair, as well as early onset obesity. From reference [387] with permission.

with hyperphagia, hyperinsulinemia and hyperglycemia, increased linear growth, and susceptibility to a wide variety of epithelial and mesenchymal tumors (reviewed in [405]). This phenotype is the result of a 170 kb deletion which removes all but the promoter and noncoding first exon of the Raly gene 5¢ to the exons of the agouti gene [406] and results in ubiquitous overexpression of agouti in all tissues examined to date [398,407]. The lethal yellow phenotype is presumably the result of agouti antagonism of the widely distributed melanocortin receptors and is consistent with the known actions of melanocortin receptors described above. An agouti-related protein (AGRP) gene was recently identified based on its homology to agouti and is expressed in the hypothalamus, adrenal medulla, and at low levels

Chapter 2

in the testis, lung, and kidney [408,409]. In contrast to the divergent expression patterns of murine and human agouti, the expression pattern of murine and human AGRP appears identical [408]. Within the hypothalamus, AGRP expression is confined to the arcuate nucleus and AGRP-immunoreactive terminals paralleled POMCimmunoreactive terminals projecting from the arcuate nucleus [55]. AGRP is a selective, nanomolar competitiveantagonist of MC3R and MC4R, clearly implicating it as the endogenous melanocortin antagonist involved in energy homeostasis [410–412]. Corticostatins are a family of related low molecular weight members of the defensin family of peptides that are competitive inhibitors of ACTH-induced steroidogenesis in the adrenal cortex, which act by blocking the ACTH receptor [413]. In rats, a dose of 8 mg/kg body weight blocks the corticosterone response to stress [414]. Corticostatins and the agouti peptides are the two known endogenous competitive inhibitors of melanocortin receptors, but do not appear to be structurally related [415].

Additional Potential Melanocortin Actions Additional physiologic roles for melanocortins have been proposed that do not clearly correlate with the known functions of specific melanocortin receptors as surmised from human or mouse mutants. A multitude of behavioral and psychological effects have been attributed to melanocortins (for review see [416]). Intracerebroventricular administration of ACTH or a-MSH elicits excessive grooming behavior [417], yawning, stretching, and penile erection [418]. The grooming, stretching, and yawning behavior, but not erectile function, may be mediated by MC4R as deduced through the use of MC4R selective antagonists [233,419,420]. Recently, in a double-blind, placebo controlled crossover study, a cyclic a-MSH analogue initiated erections in men with psychogenic erectile dysfunction [421]. Roles for the actions of melanocortins in neuromuscular development, promotion of the regeneration of crushed nerves, and CNS protection from injury have also been postulated [235,422–425]. A potential role for melanocortins in fetal growth and brain development has been demonstrated [426,427]. Data indicating a role for melanocortins as modulators of inflammation have been discussed previously. Centrally administered MSH is a potent antipyretic agent and, in an endogenous pyrogen-induced fever model in rabbits, is approximately 25,000-fold more potent than acetaminophen [428]. Physiologic roles for melanocortins in maintenance of cardiovascular homeostasis have also been proposed. Peripheral or central administration of g-MSH causes tachycardia and pressor effects [429,430]. Central administration of a-MSH results in bradycardia and depressor effects [429,430]. Melanocortins may further influence cardiovascular homeostasis through their effects on electrolyte regulation. a-MSH and g-MSH are potent natriuretics [431–434].

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SECRETION OF ACTH: PHYSIOLOGY Physiologically a number of factors interact to determine the final pattern of ACTH release, including circadian rhythms, stress, and negative feedback by glucocorticoids. These factors impact on each other in an integrated fashion to control ACTH release. The stage of development, from fetal life through puberty, and pregnancy and parturition, also impact on ACTH secretion. Finally, the immune system interacts with the HPA axis, adding another facet to the complexity of ACTH release.

Secretion Dynamics of ACTH in vivo ACTH secretion is characterized by pulsatile release of ACTH from the corticotroph in a burst-like pattern, with no interpulse secretion [435]. Fifteen-minute sampling reveals approximately 12 ACTH and cortisol pulses over a 24-hour period [436], whereas more frequent 10-minute sampling reveals 40 ACTH pulses in 24 hours [435]. Blood ACTH rises by an average of 24 pg/ml per pulse [435]. bendorphin secretion parallels the pulsatile release of ACTH [437]. Spontaneous ACTH and cortisol pulses correlate highly. There is a strong relationship between the magnitude of concomitant ACTH and cortisol pulses, particularly if a 15 minute phase delay in cortisol secretion is allowed for [435,438–440]. Not all spontaneous ACTH and cortisol pulses are concomitant: approximately 50% to 75% of spontaneous ACTH pulses are followed by a cortisol pulse, whereas approximately 60% to 90% of spontaneous cortisol pulses are preceded by an ACTH pulse [436,440]. The 24-hour pattern of ACTH pulses, but not the cortisol pattern, differs between males and females. Males have more pulses, greater mean peak ACTH amplitude, greater 24-hour ACTH secretion, and higher mean ACTH levels [438]. The sensitivity of the adrenal cortex, or the availability of ACTH to the adrenal cortex, may be greater in females. Alternatively, males and females may have different set points for cortisol feedback [438]. CRH may also be secreted in a pulsatile fashion, accounting for pulsatile ACTH secretion. When exogenous CRH is given continuously, there is a progressive desensitization of the ACTH response to CRH [441]. However, when CRH is given in a prolonged pulsatile manner, it does not desensitize the corticotroph CRH receptor, and the releasable ACTH pool is not depleted [436].

Negative Regulation of ACTH Secretion Negative feedback can be defined as long, short, or ultrashort depending on the location and nature of the hormone mediating the feedback. Long Feedback by Glucocorticoids

Long feedback refers to the effects of adrenal glucocorticoids on ACTH secretion at the pituitary and in the hypo-

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thalamus. Glucocorticoid-mediated negative feedback can be subdivided into fast and delayed, which can be further subdivided into intermediate and slow feedback. In animals, the initial fast negative effect of glucocorticoids occurs within seconds to minutes [183], involves inhibition of stimulated ACTH and CRH release, not synthesis, and occurs during the period when plasma glucocorticoid levels are increasing. Cortisol given to patients at the start of surgery attenuates the surgery-induced ACTH rise [211], and may be an example of fast feedback. Delayed feedback has two components, intermediate and slow [442]. Intermediate feedback is the component of delayed feedback that is due to inhibition of ACTH release, but not synthesis, and may be important after short durations of glucocorticoid exposure, or after noncontinuous, repeated exposures [442]. Intermediate delayed feedback develops after 45 to 120 minutes, and maximal inhibition occurs 2 to 4 hours after administration of one dose of glucocorticoids. Unlike ACTH, CRH synthesis as well as release may be affected by intermediate feedback [442]. The slow component of delayed feedback is most important after long exposures to a moderately high dose of glucocorticoid, and is a function of the total dose of glucocorticoids, the glucocorticoid level achieved, and the amount of time since the steroid was given [442]. Slow feedback occurs after more than 24 hours of exposure to glucocorticoids and can persist for days. POMC biosynthesis is inhibited, leading to inhibition of basal and stimulated ACTH secretion [442], and intracellular ACTH decreases implying decreased synthesis. In humans, negative feedback by glucocorticoids takes 30 to 60 minutes to manifest. When endogenous cortisol is suppressed with metyrapone, the effects of exogenous glucocorticoids on the morning ACTH rise, or on oCRHstimulated ACTH release, are not seen initially but appears 30 minutes after glucocorticoid administration [443]. In addition, glucocorticoids have little effect on the initial CRH-induced increase in ACTH release, but decrease CRH-induced ACTH release after 60 minutes. Cortisol given to patients at the start of surgery attenuates the surgery-induced ACTH rise [211]. Cortisol modulates the responsiveness of the pituitary. The corticotroph is dependent on CRH stimulation to maintain ACTH secretion, and glucocorticoids suppress CRH-induced ACTH secretion in vivo and in vitro. On the other hand, when endogenous cortisol levels are suppressed by metyrapone, basal ACTH and CRH-induced ACTH release are increased [444]. Glucocorticoid inhibition of ACTH secretion from the corticotroph may recover more quickly than CRH secretion from the hypothalamus [445]. Secondary adrenal insufficiency due to long-term glucocorticoid therapy may in part be due to continued suppression of hypothalamic CRH secretion 184. Adrenalectomized patients on exogenous glucocorticoid therapy have a blunted ACTH response to CRH that normalizes after several CRH boluses, suggesting that lack of stimula-

tion of the corticotroph by CRH suppresses the ACTH response [446]. On the other hand, corticotrophs of patients recovering from transsphenoidal surgery for Cushing’s disease are profoundly unresponsive to CRH, which can not be attributed solely to deficient CRH priming [447]. Glucocorticoids inhibit AVP secretion [448]. In most studies, patients with hypopituitarism or primary adrenal insufficiency are unable to maximally dilute their urine in response to a water load, and this is corrected by glucocorticoid administration [449]. However, it is unclear if the elevated AVP levels, or the lack glucocorticoids, is responsible for the inability to maximally dilute the urine in the hypocortisolemic state [448]. Glucocorticoids inhibit nitric oxide synthase, and nitric oxide is capable of stimulating the insertion of the water channel, aquaporin 2, in the luminal membrane of the renal collecting cell [450]. This may provide an explanation for why glucocorticoid deficiency is associated with decreased free water clearance. Short Feedback

Short feedback refers to the effect of pituitary ACTH to inhibit CRH release. In normal subjects, the administration of ACTH does not affect CRH levels, most likely because of the negative effects of cortisol present prior to ACTH administration [451]. However, in patients with elevated CRH levels due to Addison’s disease or hypopituitarism, ACTH decreases CRH and b-endorphin levels, suggesting that ACTH inhibits CRH secretion [451]. ACTH may act in the median eminence or in the hypothalamus to inhibit CRH release [451].

Circadian Regulation of ACTH Secretion There is an endogenous circadian rhythm to the pulsatile pattern of ACTH secretion which leads to a circadian rhythm of glucocorticoid release. The function of this circadian rhythm in cortisol secretion is not known, although one hypothesis suggests that the early morning rise in cortisol causes a delayed-phase rise in insulin resistance, which may play a role in altered glucose metabolism [452]. For instance, since the brain does not require insulin for glucose uptake, peripheral insulin resistance might cause a rise in glucose levels, leading to greater uptake in the brain. The circadian rhythm is generated in the suprachiasmatic nucleus (SCN), and the signals travel via efferent inputs to the PVN to modulate CRH release [183,453]. This circadian rhythm is due to variation in ACTH pulse amplitude, not frequency [435]. The amount of ACTH secreted per pulse varies by 3.8-fold over a 24-hour period [435]. Basal ACTH and cortisol levels parallel each other and are the highest upon awakening in the morning between 0600 hours and 0900 hours, decline through the day to intermediate levels at 1600 hours, and are lowest between 2300 hours and 0300 hours [435]. From 2300 hours to 0200 hours, there is a quiescent period of minimal secretory activity, corresponding to the nadir of ACTH and cortisol levels [454]. Secretion of

Chapter 2

ACTH and cortisol abruptly increases in the early morning [454]. The diurnal secretory pattern is similar for free and total cortisol, although the relative increase in free cortisol is about 1.5 times greater than the relative increase in total cortisol [454]. Alterations in feeding and sleep impact on cortisol secretion [455]. Cortisol briefly increases postprandially, especially after the midday meal [454]. Exercise or administration of ACTH at 1000 hours leads to a rise in cortisol and blunts the midday cortisol surge, and at 1400 hours leads to a rise in cortisol [454]. Overall, the major features of the diurnal cortisol pattern persist under conditions of complete fasting, continuous feeding, or total sleep deprivation [456]. However, the circadian rhythm of cortisol secretion fully adapts to permanent changes in environmental time and the sleep–wake pattern. This adaptation takes about 3 weeks, the limiting factor being the time it takes for the quiescent period of secretion to fully adapt [456]. The acrophase adapts much more quickly and is partially synchronized after one day and totally synchronized after 10 days [456]. There is circadian regulation of the sensitivity of the response of the adrenal cortex to ACTH [457,458]. Injection of ACTH at 0700 hours, just at the time of the endogenous cortisol peak, causes a significant increase in cortisol. The absolute increment in cortisol secretion in response to an ACTH stimulation test is greater when the test is performed at circadian nadir compared to the peak, although in one study this was true in males only [459]. The incremental increase in cortisol secretion in response to a CRH stimulation test is also greatest at night when basal levels are lowest, although the total amount of cortisol released is greatest in the morning when the basal cortisol levels are highest [460]. The basal cortisol level, not the time of day, seems to be the important factor determining the cortisol response to oCRH, and the higher the basal cortisol level, the lower the peak cortisol response to oCRH [436]. On the other hand, the maximum ACTH blood level in response to oCRH occurs at 0700 hours, the time of the minimum cortisol increment to oCRH [436]. The factors governing the circadian rhythm in ACTH release in humans are not clear. On the one hand, it may be regulated by a diurnal rhythm in CRH secretion [461]. The highest CRH levels occur at 0600 hours (7.0 pg/ml) and the lowest at 1800 hours and 2200 hours (about 1.8 pg/ml), which parallels the pattern of ACTH and cortisol secretion [462]. Serotoninergic and cholinergic pathways may play a role in the CRH circadian rhythm, and display circadian periodicity in their hypothalamic concentrations [463]. On the other hand, when the pituitary is exposed to constant levels of CRH 30-fold higher than those found in portal hypophyseal blood, such as in pregnancy [464] and during CRH infusion [441], the circadian rhythms in ACTH and cortisol persist. CRH clearly plays a role in the ACTH rhythm, as CRH-deficient mice have an absent or markedly attenuated diurnal rise in ACTH [465]. However, in these mice, constant infusion of CRH restores the ACTH

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rhythm. This indicates that changes in CRH amplitude are not necessary to drive the ACTH rhythm, but that a tonic level of CRH is required to maintain ACTH responsiveness [466] to circadian cues.

Physical Stress Regulation of ACTH Secretion Glucocorticoid release mediated by ACTH, plays a major role in the response to stress. An interaction exists between stress-mediated ACTH release, which leads to glucocorticoid secretion, and glucocorticoid-mediated negative feedback, which inhibits further ACTH and glucocorticoid release. A number of stressful stimuli lead to ACTH secretion. Most physical stressors activate the HPA axis. The magnitude of the rise in ACTH and cortisol is dependent upon many factors, including the nature of the stress, its magnitude including the rapidity of its appearance, and the time of day it is experienced. In general, stressors have a larger impact on ACTH release when they develop rapidly, are of high magnitude, and occur during the circadian nadir in ACTH release. Hypoglycemia

In humans, insulin induced hypoglycemia is associated with a increase in plasma ACTH levels [467], from a basal level of about 40 pg/ml to a peak of 250 pg/ml at 45 minutes [467]. Cortisol levels increase over twofold, from a basal level of about 11 ug/dl to a peak of about 25 ug/dl at 60 to 90 minutes [467]. Insulin-induced hypoglycemia causes a four- to fivefold greater increase in ACTH secretion than oCRH alone, and a 1.3-fold greater increase than AVP plus CRH [97,468]. CRH may play a permissive rather than a dynamic role in the ACTH response to hypoglycemia [182], whereas AVP may play a more direct role [467]. AVP levels have been shown to increase 2.5- to sevenfold at 30 to 45 minutes after the insulin. When AVP levels are raised endogenously by saline infusion or lowered to undetectable levels by waterloading, the hypoglycemia-induced AVP increase is greater after saline, even though saline blunts the hypoglycemic response to insulin [469]. Catecholamines increase in response to hypoglycemia, and may act at the hypothalamus to mediate ACTH release. Epinephrine appears to play more of a role than norepinephrine, increasing at least 13fold at 30 minutes after insulin, whereas norepinephrine increases 2.4-fold at 60 minutes. Exercise

Exercise increases ACTH and b-endorphin levels, and the response is dependent on the intensity of exercise and the level of training [470]. Exercising to exhaustion, or exercise of short duration and high intensity, increases ACTH, bendorphin, and cortisol levels. Hypercortisolism is seen in highly trained athletes, who need a higher level of oxygen consumption to stimulate ACTH release, and have elevated basal ACTH levels [471]. Physical exercise and stress both

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lead to analgesia in man [472]. Naloxone reverses exerciseinduced analgesia from certain types of pain, suggesting a role for endorphins. Dexamethasone reverses exerciseinduced analgesia from other types of pain, like dental pain, that are not reversed by naloxone, suggesting a role for ACTH, although dexamethasone also suppresses bendorphin release. Lower Body Negative Pressure and Acute Hemorrhage

Lower body negative pressure in humans simulates acute hemorrhage. Lower body negative pressure increases ACTH secretion to peak values of 60–250 pg/ml at 2 to 10 minutes after cessation of the stimulus, and the increase in ACTH is reversed by dexamethasone [473]. Hypovolemia may be the physiologically important stress. In animals, hemorrhage stimulates ACTH secretion primarily mediated by CRH and, to some degree, by AVP [182]. Hypovolemia increases portal blood CRH, AVP, epinephrine, and oxytocin, whereas hypotension, also a component of hemorrhage, induced by nitroprusside, increases portal blood CRH only. Surgical Stress

Surgery induces a large increase in plasma ACTH levels [284]. Patients undergoing surgery may have increased sensitivity of the adrenal cortex to ACTH [474,475]. Fentanyl, an opiate agonist, attenuates the ACTH response to surgery [211]. Evidence in adrenalectomized primates suggests that supraphysiological doses of glucocorticoids are not necessary for the animal to withstand surgical stress, but that a minimal level is necessary [474,476].

Psychological and Emotional Stress Psychological and emotional stress play a role in the hormonal stress response. ACTH levels are high in patients awaiting an insulin-tolerance test. During physical exercise, psychological and physical stress may act synergistically to increase b-endorphin and ACTH levels [477]. With increasing experience in performing a set level of exhaustive physical exercise, plasma ACTH and cortisol levels declined [478].

Fetal and Postnatal Regulation of ACTH Secretion In utero, maternal cortisol influences alterations of fetal heart rate, movement, and behavioral states [479]. By 35 weeks of gestation, there is a circadian rhythm in fetal behavioral states that is altered if the maternal diurnal variation in ACTH and cortisol secretion are abolished by maternal administration of triamicinolone [479]. Neonates, even when born prematurely, have an endogenous cortisol rhythm. Minimally and severely stressed neonates in the neonatal intensive care unit, born between 23 and 38 weeks of gestation, have a significant diurnal rhythm in cortisol and endorphin secretion, although ACTH levels do not vary sig-

nificantly [463]. Others have suggested that diurnal rhythmicity is a function of maturation, and is not present before 6 months of age [463,480,481].

Regulation of ACTH Secretion During Puberty Normal children between the ages of 1 year and 16 years do not differ from adults in ACTH, b-endorphin, and cortisol responses to CRH, and the responses of boys do not differ from girls [482]. Some of the other adrenal steroids and adrenal androgens demonstrate basal and stimulated variation with age. The CRH-stimulated androstenedi one to 17-hydroxyprogesterone ratio increases with sexual maturation, suggesting that the 17,20-desmolase activity increases with puberty [483]. The dehydroepiandrosterone response to CRH increases as children progress from stage 1 to stage 5 of puberty, and by stage 5 of puberty, dehydroepiandrosterone levels do not differ from adults [482].

HPA-Axis-Immune Interactions The immune system and the neuroendocrine system communicate with each other, and share a common set of structurally identical hormones and receptors [484–488]. Cells of the immune system synthesize biologically active neuroendocrine peptide hormones, immune cells contain receptors for neuroendocrine hormones, neuroendocrine hormones modulate immune function, and lymphokines modulate neuroendorine function. The neuroendocrine system and the immune system work together in the regulation of both the stress and the immune response, and the components include lymphoid cells, cholinergic and adrenergic neurons, cytokines and lymphokines, hormones and neuropeptides released by the endocrine glands and the CNS, receptors, and higher CNS activity which modulates these responses [489,490]. Effect of Immune System on HPA Axis

Immune cells, particularly monocytes, macrophages, and lymphocytes, produce cytokines involved in the immune response, and these cytokines activate the HPA axis. Bacterial-derived endotoxin and lipopolysacchride stimulate the release of interleukin-1, interleukin-6, and TNF-a, which are regulated by glucocorticoid feedback. The effects of cytokines on the HPA axis at the level of the brain, pituitary, and adrenal gland, are the best examples of immune modulation of a neuroendocrine system. Interleukin-1 is produced by stimulated macrophages and monocytes, and stimulates CRH release. Interleukin-1 also directly stimulates the adrenal cortex. Tumor necrosis factor is produced primarily by activated monocytes [491], and has many of the same biological activities and responds to many of the same immune challenges as interleukin-1. Tumor necrosis factor stimulates ACTH release, most likely at an extra-pituitary site. Interleukin-6 has actions similar to interleukin-1 and

Chapter 2

TNF, and stimulates the HPA axis [202,491]. Interleukin-2 is synthesized by T cells after an antigenic challenge, and increases ACTH and cortisol levels in humans, although this may be indirect and due to the stress response generated by fever and chills. Gamma-interferon causes an increase in steroid production by adrenal cells. As discussed above, leukemia-inhibitory factor (LIF) may be a major activator of POMC gene transcription and ACTH release following immune or inflammatory stimulation. Effect of HPA Axis on Immune System

Stress suppresses immune function. Following an infection or most immunization procedures, and the presence of bacterial endotoxin, a stress-like response of the pituitary occurs leading to the release of ACTH and cortisol which tends to suppress the immune system. However, in humans, there is no clear evidence that the rise in cortisol following an immune or inflammatory stimulus plays a significant role in modulating the subsequent immune response. In fact, prior to the treatment of patients with primary adrenal insufficiency, there were no substantive reports that these patients suffered from overactivity of their immune systems. Receptors for CRH, ACTH, and glucocorticoids most likely mediate the effect of these hormones on the immune system. ACTH receptors on human peripheral monocytes have been characterized, and glucocorticoid receptors are present on human lymphocytes. Glucocorticoids inhibit many aspects of immune function, establishing a negative feedback loop between the immune and neuroendocrine systems. Glucocortiocoids block lymphocyte activation, block the production and action of interleukin-2, interleukin-1, gamma-interferon, TNF, and prostaglandins, and interfer with the interaction of certain effector molecules with target cells. HPA Axis Within Immune System

Cells of the immune system produce CRH, ACTH and endorphins. Human peripheral mononuclear leukocytes synthesize three molecular forms of immunoreactiveACTH, and lymphocytes produce b-endorphins. Mouse spleen macrophages and virally infected mouse splenocytes contain b-endorphin and POMC mRNA [492–494]. CRH mRNA and peptide are found in monocytes in acute inflammatory reactions [495] and in T cell lymphocytes [496]. Releasing hormones and cytokines interact to stimulate ACTH production from immune cells. CRH and AVP stimulate a dramatic increase in biologically active ACTH and b-endorphin from human peripheral leukocytes in some studies. Unstimulated leukocytes produce little ACTH. In humans, only B lymphocytes secrete b-endorphin in response to CRH and AVP, and monocyte-secreted interleukin-1 mediates the effect. A feedback loop exists between the immune and neuroendocrine systems [490]. Cytokines released by immune cells stimulate secretion of ACTH and glucocorticoids which are active in the fight

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against infection. Subsequently, these hormones suppress the further synthesis of cytokines. SECRETION OF ACTH: PATHOPHYSIOLOGY Abnormal secretion of ACTH occurs in ACTH-secreting pituitary adenomas responsible for Cushing’s disease or Nelson’s syndrome, and in Cushing’s syndrome due to an ectopic pituitary adenoma or an ectopic ACTH-secreting tumor. Decreased or absent ACTH secretion due to genetic or autoimmune causes has been discussed in the above section on melanocortin actions.

Pituitary Adenomas ACTH immunoreactivity in an abnormal pituitary is usually due to a single functional adenoma, but can be associated with nodular hyperplasia or a silent cortiocotropic adenoma. Diffuse or nodular corticotroph hyperplasia, which could result from an ectopic or hypothalmic CRH-producing tumor, or Addison’s disease, is a rare cause of a pituitary adenoma or ACTH hyperfunction. Pituitary adenomas constitute about 15% of intracranial tumors [3]. Approximately 56% of pituitary adenomas are active, and about one third of active pituitary adenomas produce ACTH. Ten to 15% of pituitary adenomas are pleurihormonal, some of which secrete ACTH. When discovered, ACTH-producing adenomas are often functional, small, highly vascular, and prone to hemorrhage [497]. Corticotroph adenomas tend to be located in the central portion of the adenohypophysis, in the “mucoid wedge,” and form micronodular aggregates [497]. Invasive adenomas are more frequently found among undifferentiated, extermely laterally localized, or large adenomas, and recur more frequently than non-invasive tumors. There is no correlation between the size of an adenoma and the cortisol level or rate of recurrence [498]. The majority of adenomas responsible for Cushing’s disease or Nelson’s syndrome are microadenomas. ACTH-producing adenomas are usually monoclonal, but may be polyclonal [499]. Pleurihormonal adenomas are usually polyclonal [499]. The biochemical structure and ultrastructure of ACTHsecreting pituitary adenoma cells differ from nonadenomatous pituitary cells [500]. The cells may look normal, but be increased in number [497]. Cells are oval to polygonal in shape, with eccentric spherical nuclei and well-developed rough endoplasmic reticulim. Crooke’s hyaline is characteristic of ACTH-producing tumors [501], is associated with either endogenous or exogenous hypercortisolism, and is due to massive accumulation of intermediate cytoplasmic filaments that are normally present in small numbers [3]. Clinically, an increased number of Crooke’s cells is correlated with a longer postoperative replacement dose of cortisol requirement [498]. Pituitary adenomas may produce products in addition to ACTH. Some ACTH-producing corticotroph adenomas contain a form of gastrin that is smaller than gastrin found

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in the normal adenophypophysis. 7B2 is a secretory granule-associated protein, which may be involved in proconvertase activtion [502], which is sometimes secreted by ACTH-producing tumors, it is secreted in the highest levels from nonfunctional pituitary tumor. A variant of Cushing’s disease is due to ACTHproducing pituitary adenomas occurring ectopically [5], and have been described to arise in the mucosa of the sphenoid sinus [5,503], within a benign cystic ovarian teratoma [504], and intrahemispherically in a 6-year-old boy [505]. Rarely, both Cushing’s disease and Nelson’s syndrome have been preceded by generalized glucocorticoid resistance due to a mutation in the glucocorticoid receptor [506,507]. In these cases, the high rate of ACTH secretion, stimulated by the generalized glucocorticoid resistance, in some way led to adenoma formation, perhaps following a second oncogenic transformation. Pituitary carcinomas are malignant pituitary tumors associated with extracranial metastases, including liver and lung [508]. Pituitary carcinomas cannot be histologically differentiated from adenomas [3]. A large number of pituitary carcinomas produce Cushing’s disease [3]. In patients with Cushing’s disease due to a pituitary carcinoma, the primary tumor and metastases stain immunochemically for ACTH, b-LPH, b-endorphin, and a-MSH [508], and production of both CRH and ACTH from a pituitary carcinoma has been described [509].

Most nonpituitary POMC-secreting tumors do not produce CRH [515]. However, some patients with ectopic ACTH syndrome due to lung cancer have tumors that produce ACTH and CRH, and secrete ACTH in response to CRH [516]. Bronchial carcinoid tumors may contain CRH and ACTH, and be associated with high plasma ACTH and CRH levels and Cushing’s syndrome, or with normal plasma CRH levels. Cushing’s syndrome has been associated with ectopic ACTH secretion from a unilateral adrenal pheochromocytoma [517], and from bilateral pheochromocytomas in a case of multiple endocrine neoplasia type 2A [518]. A number of other ectopic tumors have been described that secrete ACTH, including an adenoid cystic carcinoma of the lung [519], a renal cell carcinoma [520], a neuroendocrine tumor of the nasal roof [521], and an ACTHproducing tumor metastatic to the liver [522]. In general, POMC mRNA from nonpituitary tumors responsible for the ectopic ACTH syndrome is identical to normal and to POMC mRNA from pituitary tumors [523]. However, some tumors contain a larger POMC mRNA species that is increased in amount from 0.3% of the overall POMC mRNA in normal pituitaries, to up to 35 to 40% in the tumor, and is transcribed from an alternative upstream promoter [523]. Pancreatic islet cell tumors responsible for Cushing’s syndrome have been demonstrated to contain ACTH and b-endorphin [524], and to express POMC mRNA [524].

Ectopic ACTH-secreting Tumors Several neuroendocrine neoplasms occur in the bronchopulmonary tract, including small-cell neuroendocrine carcinomas, carcinoids, well-differentiated neuroendocrine carcinomas, and intermediate-cell neuroendocrine carcinomas [510]. These neoplasms express neuroendocrine markers including chromogranins and synaptophysin [510]. Thirtyfour percent of small-cell carcinomas of the lung show immunoreactivity to one or more peptide hormones [511], and patients with peptide-positive small-cell lung carcinomas have a shorter mean survival than in patients with nonreactive tumors. ACTH is by far the most common hormone present, and is seen in 24% of small-cell carcinomas [511]. However, 56% of small-cell lung cancer cell lines secrete significant concentrations of ACTH precursors, with little, if any, processing to ACTH. Oat-cell lung carcinomas may also produce ACTH, leading to Cushing’s syndrome [512]. Carcinoid tumors often produce more than one hormone, and can be responsible for the ectopic ACTH syndrome [513]. Bronchial carcinoids occasionally contain ACTH and related opioid peptides, which does not alter the overall favorable prognosis of these tumors. Recently, Cushing’s syndrome caused by ACTH secretion by pulmonary tumorlets has been described [514]. Upon radiologic imaging, such tumorlets, which may be over 100 in number, present a very unusual appearance.

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187 Jones MT, Hillhouse EW, Burden J. Effect on various putative neurotransmitters on the secretion of corticotrophin-releasing hormone from the rat hypothalamus in vitro – a model of the neurotransmitters involved. J Endocrinol 1976;69:1–10. 188 Rivier C, Vale W. Effects of corticotropin-releasing factor, neurohypophyseal peptides, and catecholamines on pituitary function. Fed Proc 1985;44: 189–195. 189 al-Damluji S. Adrenergic mechanisms in the control of corticotrophin secretion. J Endocrinol 1988;119:5–14. 190 Watanabe T, Oki Y, Orth DN. Kinetic actions and interactions of arginine vasopressin, angiotensin-II, and oxytocin on adrenocorticotropin secretion by rat anterior pituitary cells in the microperifusion system. Endocrinology 1989; 125:1921–1931. 191 Schoenenberg P, Gaillard RC, Kehrer P et al. cAMP-dependent ACTH secretagogues facilitate corticotropin releasing activity of angiotensin II on rat anterior pituitary cells in vitro. Acta Endocrinol (Copenh) 1987;114:118–123. 192 McCann SM, Rettori V, Milenkovic L et al. Role of monokines in control of anterior pituitary hormone release. Adv Exp Med Biol 1990;274:315–329. 193 Berkenbosch F, van Oers J, del Rey A et al. Corticotropin-releasing factorproducing neurons in the rat activated by interleukin-1. Science 1987;238: 524–526. 194 Sapolsky R, Rivier C, Yamamoto G et al. Interleukin-1 stimulates the secretion of hypothalamic corticotropin-releasing factor. Science 1987;238:522–524. 195 Matta S, Singh J, Newton R et al. The adrenocorticotropin response to interleukin-1 beta instilled into the rat median eminence depends on the local release of catecholamines. Endocrinology 1990;127:2175–2182. 196 Michie HR, Majzoub JA, ODwyer ST et al. Both cyclooxygenase-dependent and cyclooxygenase-independent pathways mediate the neuroendocrine response in humans. Surgery 1990;108:254–259. 197 Michie HR, Spriggs DR, Manogue KR et al. Tumor necrosis factor and endotoxin induce similar metabolic responses in human beings. Surgery 1988;104:280–286. 198 Gaillard RC, Turnill D, Sappino P et al. Tumor necrosis factor alpha inhibits the hormonal response of the pituitary gland to hypothalamic releasing factors. Endocrinology 1990;127:101–106. 199 Sharp BM, Matta SG, Peterson PK et al. Tumor necrosis factor-alpha is a potent ACTH secretagogue: comparison to interleukin-1 beta. Endocrinology 1989;124:3131–3133. 200 Michie HR, Manogue KR, Spriggs DR et al. Detection of circulating tumor necrosis factor after endotoxin administration. N Engl J Med 1988;318: 1481–1486. 201 Yasuda N, Greer MA. Evidence that the hypothalamus mediates endotoxin stimulation of adrenocorticotropic hormone secretion. Endocrinology 1978; 102:947–953. 202 Naitoh Y, Fukata J, Tominaga T et al. Interleukin-6 stimulates the secretion of adrenocorticotropic hormone in conscious, freely-moving rats. Biochem Biophys Res Commun 1988;155:1459–1463. 203 Navarra P, Tsagarakis S, Faria MS et al. Interleukins-1 and -6 stimulate the release of corticotropin-releasing hormone-41 from rat hypothalamus in vitro via the eicosanoid cyclooxygenase pathway. Endocrinology 1991;128:37–44. 204 Bochicchio D, Ambrosi B, Faglia G. Loperamide, an opiate analog, differently modifies the adrenocorticotropin responses to corticotropin-releasing hormone and lysine vasopressin in patients with Addison’s disease. Neuroendocrinology 1988;48:611–614. 205 Atkinson RL. Endocrine and metabolic effects of opiate antagonists. J Clin Psychiatry 1984;45:20–24. 206 Gaillard RC, Grossman A, Smith R et al. The effects of a met-enkephalin analogue on ACTH, beta-LPH, beta-endorphin and MET-enkephalin in patients with adrenocortical disease. Clin Endocrinol (Oxf ) 1981;14:471–478. 207 Blankstein J, Reyes FI, Winter JS et al. Effects of naloxone upon prolactin and costisol in normal women. Proc Soc Exp Biol Med 1980;164:363–365. 208 Grossman A, Gaillard RC, McCartney P et al. Opiate modulation of the pituitary–adrenal axis: effects of stress and circadian rhythm. Clin Endocrinol (Oxf) 1982;17:279–286. 209 Morley JE, Baranetsky NG, Wingert TD et al. Endocrine effects of naloxoneinduced opiate receptor blockade. J Clin Endocrinol Metab 1980;50:251–257. 210 Volavka J, Cho D, Mallya A et al. Naloxone increases ACTH and cortisol levels in man [letter]. N Engl J Med 1979;300:1056–1057. 211 Raff H, Flemma RJ, Findling JW. Fast cortisol-induced inhibition of the adrenocorticotropin response to surgery in humans. J Clin Endocrinol Metab 1988;67:1146–1148. 212 Spiler IJ, Molitch ME. Lack of modulation of pituitary hormone stress response by neural pathways involving opiate receptors. J Clin Endocrinol Metab 1980;50:516–520.

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264 Hartmeyer M, Scholzen T, Becher E et al. Human dermal microvascular endothelial cells express the melanocortin receptor type 1 and produce increased levels of IL-8 upon stimulation with alpha-melanocyte-stimulating hormone. J Immunol 1997;159:1930–1937. 265 Haynes RC. The activation of adrenal phosphorylase by the adrenocorticotropic hormone. J Biol Chem 1958;233:1220–1222. 266 Lefkowitz RJ, Roth J, Pricer W et al. ACTH receptors in the adrenal: specific binding of ACTH-125I and its relation to adenyl cyclase. Proc Natl Acad Sci USA 1970;65:745–752. 267 Catalano RD, Stuve L, Ramachandran J. Characterization of corticotropin receptors in human adrenocortical cells. J Clin Endocrinol Metab 1986;62: 300–304. 268 Buckley DI, Ramachandran J. Characterization of corticotropin receptors on adrenocortical cells. Proc Natl Acad Sci USA 1981;78:7431–7435. 269 Dallman MF, Akana SF, Jacobson L et al. Characterization of corticosterone feedback regulation of ACTH secretion. Ann NY Acad Sci 1987;512:402–14. 270 Weber A, Kapas S, Hinson J et al. Functional characterization of the cloned human ACTH receptor: impaired responsiveness of a mutant receptor in familial glucocorticoid deficiency. Biochem Biophys Res Commun 1993;197: 172–178. 271 Naville D, Penhoat A, Barjhoux L et al. Characterization of the human ACTH receptor gene and in vitro expression. Endocr Res 1996;22:337–348. 272 Elias LL, Huebner A, Pullinger GD et al. Functional characterization of naturally occurring mutations of the human adrenocorticotropin receptor: poor correlation of phenotype and genotype. J Clin Endocrinol Metab 1999;84: 2766–2770. 273 Brown MS, Kovanen PT, Goldstein JL. Receptor-mediated uptake of lipoprotein-cholesterol and its utilization for steroid synthesis in the adrenal cortex. Recent Prog Horm Res 1979;35:215–257. 274 Pedersen RC, Brownie AC. Adrenocortical response to corticotropin is potentiated by part of the amino-terminal region of procorticotropin/endorphin. Proc Natl Acad Sci USA 1980;77:2239–2243. 275 Jefcoate CR, Simpson ER, Boyd GS. Spectral properties of rat adrenalmitochondrial cytochrome P-450. Eur J Biochem 1974;42:539–551. 276 Privalle CT, Crivello JF, Jefcoate CR. Regulation of intramitochondrial cholesterol transfer to side-chain cleavage cytochrome P-450 in rat adrenal gland. Proc Natl Acad Sci USA 1983;80:702–706. 277 Papadopoulos V, Brown AS, Hall PF. Calcium-calmodulin-dependent phosphorylation of cytoskeletal proteins from adrenal cells. Mol Cell Endocrinol 1990;74:109–123. 278 Miller WL, Strauss JF. Molecular pathology and mechanism of action of the steroidogenic acute regulatory protein, StAR. J Steroid Biochem Mol Biol 2001; 69:131–141. 279 Simpson ER. Cholesterol side-chain cleavage, cytochrone P450, and the control of steroidogenesis. Mol Cell Endocrinol 1979;13:213–227. 280 Yago N, Ichii S. Submitochondrial distribution of components of the steroid 11 beta-hydroxylase and cholesterol sidechain-cleaving enzyme systems in hog adrenal cortex. J Biochem (Tokyo) 1969;65:215–224. 281 Churchill PF, Kimura T. Topological studies of cytochromes P-450scc and P-45011 beta in bovine adrenocortical inner mitochondrial membranes. Effects of controlled tryptic digestion. J Biol Chem 1979;254:10443–10448. 282 Orme-Johnson NR. Distinctive properties of adrenal cortex mitochondria. Biochim Biophys Acta 1990;1020:213–231. 283 Miller WL. Molecular biology of steroid hormone synthesis. [Review]. Endocrine Reviews 1988;9:295–318. 284 Saez JM, Begeot M, Durand P. [ACTH receptors]. Ann Endocrinol (Paris) 1989; 50:409–417. 285 Tepperman J, Engel FL, Long CNH. A review of adrenal cortical hypertrophy. Endocrinol 1943;32:373–402. 286 Smith PE. Hypophysectomy and a replacement therapy in the rat. Amer J Anta 1930;45:205–273. 287 Ney RL, Dexter RN, Davis WW et al. A study of mechanisms by which adrenocorticotropic hormone maintains adrenal steroidogenic responsiveness. J Clin Invest 1967;46:1916–1924. 288 Ingle DJ. The functional interrelationship of anterior pituitary and the adrenal cortex. Ann Intern Med 1951;35:652–672. 289 Wyllie AH, Kerr JF, Macaskill IA et al. Adrenocortical cell deletion: the role of ACTH. J Pathol 1973;111:85–94. 290 Farese RV, Reddy WJ. Observations on the interrelations between adrenal protein, RNA and DNA during prolonged ACTH administration. Biocem Biophys Acta 1963;76:145–148. 291 Imai T, Seo H, Murata Y et al. Adrenocorticotropin increases expression of c-fos and beta-actin genes in the rat adrenals. Endocrinology 1990;127: 1742–1747.

292 Gospodarowicz D, Ill CR, Hornsby PJ et al. Control of bovine adrenal cortical cell proliferation by fibroblast growth factor. Lack of effect of epidermal growth factor. Endocrinology 1977;100:1080–1089. 293 Masui H, Garren LD. Inhibition of replication in functional mouse adrenal tumor cells by adrenocorticotropic hormone mediated by adenosine 3¢:5¢cyclic monophosphate. Proc Natl Acad Sci USA 1971;68:3206–3210. 294 Ramachandran J, Suyama AT. Inhibition of replication of normal adrenocortical cells in culture by adrenocorticotropin. Proc Natl Acad Sci USA 1975;72:113–117. 295 Rao AJ, Long JA, Ramachandran J. Effects of antiserum to adrenocorticotropin on adrenal growth and function. Endocrinology 1978;102:371–378. 296 Dallman MF, Engeland WC, Holzwarth MA et al. Adrenocorticotropin inhibits compensatory adrenal growth after unilateral adrenalectomy. Endocrinology 1980;107:1397–1404. 297 Estivariz FE, Iturriza F, McLean C et al. Stimulation of adrenal mitogenesis by N-terminal proopiocortin peptides. Nature 1982;297:419–422. 298 Estivariz FE, Morano MI, Carino M et al. Adrenal regeneration in the rat is mediated by mitogenic N-terminal pro-opiomelanocortin peptides generated by changes in precursor processing in the anterior pituitary. J Endocrinol 1988;116:207–216. 299 Engeland WC, Shinsako J, Dallman MF. Corticosteroids and ACTH are not required for compensatory adrenal growth. Am J Physiol 1975;229: 1461–1464. 300 Dallman MF, Engelmand WC, Shinsako J. Compensatory adrenal growth: a neurally mediated reflex. Am J Physiol 1976;231:408–414. 301 Albright F. Osteoporosis. Annals of Internal Medicine 1947;27:861–882. 302 Ishihara F, Komatsu M, Yamada T et al. Role of dehydroepiandrosterone and dehydroepiandrosterone sulfate for the maintenance of axillary hair in women. Horm Metab Res 1993;25:34–36. 303 Stewart ME, Downing DT, Cook JS et al. Sebaceous gland activity and serum dehydroepiandrosterone sulfate levels in boys and girls. Archives of Dermatology 1992;128:1345–1348. 304 Rosenfield RL, Grossman BJ, Ozoa N. Plasma 17-ketosteroids and testosterone in prepubertal children before and after ACTH administration. J Clin Endocrinol Metab 1971;33:249–253. 305 Rosenfield RL. Plasma 17-ketosteroids and 17-beta hydroxysteroids in girls with premature development of sexual hair. J Pediatr 1971;79: 260–266. 306 August GP, Hung W, Mayes DM. Plasma androgens in premature pubarche: value of 17 alpha-hydroxyprogesterone in differentiation from congenital adrenal hyperplasia. J Pediatr 1975;87:246–249. 307 Korth-Schutz S, Levine LS, New MI. Serum androgens in normal prepubertal and pubertal children and in children with precocious adrenarche. J Clin Endocrinol Metab 1976;42:117–124. 308 Warne GL, Carter JN, Faiman C et al. Hormonal changes in girls with precocious adrenarche: a possible role for estradiol or prolactin. J Pediatr 1978;92:743–747. 309 Sizonenko PC, Paunier L. Hormonal changes in puberty III: Correlation of plasma dehydroepiandrosterone, testosterone, FSH, and LH with stages of puberty and bone age in normal boys and girls and in patients with Addison’s disease or hypogonadism or with premature or late adrenarche. J Clin Endocrinol Metab 1975;41:894–904. 310 Parker LN, Odell WD. Control of adrenal androgen secretion. Endocr Rev 1980;1:392–410. 311 Vaitukaitis JL, Dale SL, Melby JC. Role of ACTH in the secretion of free dehydroepiandrosterone and its sulfate ester in man. J Clin Endocrinol Metab 1969;29:1443–1447. 312 Rosenfeld RS, Hellman L, Roffwarg H et al. Dehydroisoandrosterone is secreted episodically and synchronously with cortisol by normal man. J Clin Endocrinol Metab 1971;33:87–92. 313 Irvine WJ, Toft AD, Wilson KS et al. The effect of synthetic corticotropin analogues on adrenocortical, anterior pituitary and testicular function. J Clin Endocrinol Metab 1974;39:522–529. 314 Migeon C, Keller A, Lawrence B et al. DHA and androsterone levels in human plasma. J Clin Endocrinol Metab 1957;17:1051–1062. 315 Parker LN, Sack J, Fisher DA et al. The adrenarche: prolactin, gonadotropins, adrenal androgens, and cortisol. J Clin Endocrinol Metab 1978;46:396–401. 316 Linder BL, Esteban NV, Yergey AL et al. Cortisol production rate in childhood and adolescence [see comments]. J Pediatr 1990;117:892–896. 317 Cutler Jr GB, Glenn M, Bush M et al. Adrenarche: a survey of rodents, domestic animals, and primates. Endocrinology 1978;103:2112–2118. 318 Shepard TH, Landing BH, Mason DG. Familial Addison’s disease. Case reports of two sisters with corticoid deficiency unassociated with hypoaldosteronism. Am J Dis Child 1959;97:154–162.

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398 Bultman SJ, Michaud EJ, Woychik RP. Molecular characterization of the mouse agouti locus. Cell 1992;71:1195–1204. 399 Kiefer LL, Veal JM, Mountjoy KG et al. Melanocortin receptor binding determinants in the agouti protein. Biochemistry 1998;37:991–997. 400 Willard DH, Bodnar W, Harris C et al. Agouti structure and function: characterization of a potent alpha-melanocyte stimulating hormone receptor antagonist. Biochemistry 1995;34:12341–12346. 401 Kwon HY, Bultman SJ, Loffler C et al. Molecular structure and chromosomal mapping of the human homolog of the agouti gene. Proc Natl Acad Sci USA 1994;91:9760–9764. 402 Wilson BD, Ollmann MM, Kang L et al. Structure and function of ASP, the human homolog of the mouse agouti gene. Hum Mol Genet 1995;4:223–230. 403 Yang YK, Ollmann MM, Wilson BD et al. Effects of recombinant agoutisignaling protein on melanocortin action. Mol Endocrinol 1997;11:274–280. 404 Kiefer LL, Ittoop OR, Bunce K et al. Mutations in the carboxyl terminus of the agouti protein decrease agouti inhibition of ligand binding to the melanocortin receptors. Biochemistry 1997;36:2084–2090. 405 Yen TT, Gill AM, Frigeri LG et al. Obesity, diabetes, and neoplasia in yellow A(vy)/-mice: ectopic expression of the agouti gene. Faseb J 1994;8:479–488. 406 Michaud EJ, Bultman SJ, Klebig ML et al. A molecular model for the genetic and phenotypic characteristics of the mouse lethal yellow (Ay) mutation. Proc Natl Acad Sci USA 1994;91:2562–2566. 407 Miller MW, Duhl DM, Vrieling H et al. Cloning of the mouse agouti gene predicts a secreted protein ubiquitously expressed in mice carrying the lethal yellow mutation. Genes Dev 1993;7:454–467. 408 Shutter JR, Graham M, Kinsey AC et al. Hypothalamic expression of ART, a novel gene related to agouti, is up-regulated in obese and diabetic mutant mice. Genes Dev 1997;11:593–602. 409 Ollmann MM, Wilson BD, Yang YK et al. Antagonism of central melanocortin receptors in vitro and in vivo by agouti-related protein [published erratum appears in Science 1998 Sep 11;281(5383):1615]. Science 1997;278:135–138. 410 Rosenfeld RD, Zeni L, Welcher AA et al. Biochemical, biophysical, and pharmacological characterization of bacterially expressed human agouti-related protein. Biochemistry 1998;37:16041–16052. 411 Yang YK, Thompson DA, Dickinson CJ et al. Characterization of Agoutirelated protein binding to melanocortin receptors. Mol Endocrinol 1999; 13:148–155. 412 Fong TM, Mao C, MacNeil T et al. ART (protein product of agouti-related transcript) as an antagonist of MC-3 and MC-4 receptors. Biochem Biophys Res Commun 1997;237:629–631. 413 Zhu Q, Bateman A, Singh A et al. Isolation and biological activity of corticostatic peptides (anti-ACTH). Endocr Res 1989;15:129–149. 414 Cervini LA, Gray WR, Kaiser R et al. Rat corticostatin R4: synthesis, disulfide bridge assignment, and in vivo activity. Peptides 1995;16:837–842. 415 Tatro JB. Receptor biology of the melanocortins, a family of neuroimmunomodulatory peptides. [Review] [224 refs]. Neuroimmunomodulation 1996;3:259–284. 416 de Wied D, Jolles J. Neuropeptides derived from pro-opiocortin: behavioral, physiological, and neurochemical effects. Physiol Rev 1982;62:976–1059. 417 Dunn AJ. Studies on the neurochemical mechanisms and significance of ACTH-induced grooming. Ann NY Acad Sci 1988;525:150–168. 418 Bertolini A, Gessa GA, Ferrari W. Penile erection and ejaculation: a central effect of ACTH-like peptides in mammals. In: Sandler M, Gessa GA. eds. Sexual Behavior, Pharmacology and Biochemistry. NY: Raven Press, 1975: 247–257. 419 Vergoni AV, Bertolini A, Mutulis F et al. Differential influence of a selective melanocortin MC4 receptor antagonist (HS014) on melanocortin-induced behavioral effects in rats. Eur J Pharmacol 1998;362:95–101. 420 Argiolas A, Melis MR, Murgia S et al. ACTH- and alpha-MSH-induced grooming, stretching, yawning and penile erection in male rats: site of action in the brain and role of melanocortin receptors. Brain Res Bull 2000;51: 425–431. 421 Wessells H, Fuciarelli K, Hansen J et al. Synthetic melanotropic peptide initiates erections in men with psychogenic erectile dysfunction: double-blind, placebo controlled crossover study. J Urol 1998;160:389–393. 422 Strand FL, Stoboy H, Cayer A. A possible direct action of ACTH on nerve and muscle. Neuroendocrinology 1973;13:1–20. 423 Strand FL, Kung TT. ACTH accelerates recovery of neuromuscular function following crushing of peripheral nerve. Peptides 1980;1:135–138. 424 Strand FL, Rose KJ, King JA et al. ACTH modulation of nerve development and regeneration. Prog Neurobiol 1989;33:45–85. 425 Gispen WH, Verhaagen J, Bar D. ACTH/MSH-derived peptides and peripheral nerve plasticity: neuropathies, neuroprotection and repair. Prog Brain Res 1994;100:223–229.

Chapter 2 426 Swaab DF, Visser M, Tilders FJ. Stimulation of intra-uterine growth in rat by alpha-melanocyte-stimulating hormone. J Endocrinol 1976;70:445–455. 427 Swaab DF, Martin JT. Functions of alpha-melanotropin and other opiomelanocortin peptides in labour, intrauterine growth and brain development. Ciba Found Symp 1981;81:196–217. 428 Murphy MT, Richards DB, Lipton JM. Antipyretic potency of centrally administered alpha-melanocyte stimulating hormone. Science 1983;221: 192–193. 429 Klein MC, Hutchins PM, Lymangrover JR et al. Pressor and cardioaccelerator effects of gamma MSH and related peptides. Life Sci 1985;36:769–775. 430 Li SJ, Varga K, Archer P et al. Melanocortin antagonists define two distinct pathways of cardiovascular control by alpha- and gamma-melanocytestimulating hormones. J Neurosci 1996;16:5182–5188. 431 Lymangrover JR, Buckalew VM, Harris J et al. Gamma-2MSH is natriuretic in the rat. Endocrinology 1985;116:1227–1229. 432 Orias R, McCann SM. Natriuretic effect of alpha melanocyte stimulating hormone (-MSH) in hypophysectomized or adrenalectomized rats. Proc Soc Exp Biol Med 1972;139:872–876. 433 Orias R, McCann SM. Natriuresis induced by alpha and beta melanocyte stimulating hormone (MSH) in rats. Endocrinology 1972;90:700–706. 434 Chen XW, Ying WZ, Valentin JP et al. Mechanism of the natriuretic action of gamma-melanocyte-stimulating hormone. Am J Physiol 1997;272: R1946–R1953. 435 Veldhuis JD, Iranmanesh A, Johnson ML et al. Amplitude, but not frequency, modulation of adrenocorticotropin secretory bursts gives rise to the nyctohemeral rhythm of the corticotropic axis in man. J Clin Endocrinol Metab 1990;71:452–463. 436 Desir D, Van Cauter E, Beyloos M et al. Prolonged pulsatile administration of ovine corticotropin-releasing hormone in normal man. J Clin Endocrinol Metab 1986;63:1292–1299. 437 Veldhuis JD, Iranmanesh A, Johnson ML et al. Twenty-four-hour rhythms in plasma concentrations of adenohypophyseal hormones are generated by distinct amplitude and/or frequency modulation of underlying pituitary secretory bursts. J Clin Endocrinol Metab 1990;71:1616–1623. 438 Horrocks PM, Jones AF, Ratcliffe WA et al. Patterns of ACTH and cortisol pulsatility over twenty-four hours in normal males and females. Clin Endocrinol (Oxf ) 1990;32:127–134. 439 Iranmanesh A, Lizarralde G, Johnson ML et al. Circadian, ultradian, and episodic release of beta-endorphin in men, and its temporal coupling with cortisol. J Clin Endocrinol Metab 1989;68:1019–1026. 440 Krishnan KR, Ritchie JC, Saunders W et al. Nocturnal and early morning secretion of ACTH and cortisol in humans. Biol Psychiatry 1990;28: 47–57. 441 Schulte HM, Chrousos GP, Gold PW et al. Continuous administration of synthetic ovine corticotropin-releasing factor in man. Physiological and pathophysiological implications. J Clin Invest 1985;75:1781–1785. 442 Keller Wood ME, Dallman MF. Corticosteroid inhibition of ACTH secretion. Endocr Rev 1984;5:1–24. 443 DeBold CR, Jackson RV, Kamilaris TC et al. Effects of ovine corticotropinreleasing hormone on adrenocorticotropin secretion in the absence of glucocorticoid feedback inhibition in man. J Clin Endocrinol Metab 1989;68:431–437. 444 Totani Y, Niinomi M, Takatsuki K et al. Effect of metyrapone pretreatment on adrenocorticotropin secretion induced by corticotropin-releasing hormone in normal subjects and patients with Cushing’s disease. J Clin Endocrinol Metab 1990;70:798–803. 445 Watson AC, Rosenfield RL, Fang VS. Recovery from glucocorticoid inhibition of the responses to corticotrophin-releasing hormone. Clin Endocrinol (Oxf ) 1988;28:471–477. 446 Schopohl J, Hauer A, Kaliebe T et al. Repetitive and continuous administration of human corticotropin releasing factor to human subjects. Acta Endocrinol (Copenh) 1986;112:157–165. 447 Avgerinos PC, Nieman LK, Oldfield EH et al. The effect of pulsatile human corticotropin-releasing hormone administration on the adrenal insufficiency that follows cure of Cushing’s disease. J Clin Endocrinol Metab 1989;68: 912–916. 448 Raff H. Glucocorticoid inhibition of neurohypophysial vasopressin secretion. Am J Physiol 1987;252:R635–R644. 449 Oelkers W. Hyponatremia and inappropriate secretion of vasopressin (antidiuretic hormone) in patients with hypopituitarism [see comments]. N Engl J Med 1989;321:492–496. 450 Bouley R, Breton S, Sun T et al. Nitric oxide and atrial natriuretic factor stimulate cGMP-dependent membrane insertion of aquaporin 2 in renal epithelial cells. J Clin Invest 2000;106:115–126.

Adrenocorticotropin

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451 Suda T, Tomori N, Yajima F et al. A short negative feedback mechanism regulating corticotropin-releasing hormone release. J Clin Endocrinol Metab 1987;64:909–913. 452 Van Cauter E, Polonsky K, Scheen AJ. Role of circadian rhythm and sleep in human glucose regulation. Endocr Rev 1997;18:716–738. 453 Moore RY, Eichler VB. Loss of a circadian adrenal corticosterone rhythm following suprachiasmatic lesions in the rat. Brain Res 1972;42:201–206. 454 Follenius M, Brandenberger G. Plasma free cortisol during secretory episodes. J Clin Endocrinol Metab 1986;62:609–612. 455 Dallman MF, Strack AM, Akana SF et al. Feast and famine: critical role of glucocorticoids with insulin in daily energy flow. [Review] [26 refs.] Front Neuroendocrinol 1993;14:303–347. 456 Van Cauter E, Refetoff S. Multifactorial control of the 24-hour secretory profiles of pituitary hormones. J Endocrinol Invest 1985;8:381–391. 457 Dallman MF, Engeland WC, Rose JC et al. Nycthemeral rhythm in adrenal responsiveness to ACTH. Am J Physiol 1978;235:R210–R218. 458 Dijkstra I, Binnekade R, Tilders FJ. Diurnal variation in resting levels of corticosterone is not mediated by variation in adrenal responsiveness to adrenocorticotropin but involves splanchnic nerve integrity. Endocrinology 1996;137:540–547. 459 Iannotta F, Magnoli L, Visconti G et al. Differences in cortisol, aldosterone and testosterone responses to ACTH 1–17 administered at two different times of day. Chronobiologia 1987;14:38–46. 460 Sensi S, Capani F, De Remigis P et al. Circadian time structure of pituitary and adrenal response to CRF, TRH and LHRH. Prog Clin Biol Res 1990;341:535–542. 461 Suda T, Tomori N, Yajima F et al. Immunoreactive corticotropin-releasing factor in human plasma. J Clin Invest 1985;76:2026–2029. 462 Watabe T, Tanaka K, Kumagae M et al. Diurnal rhythm of plasma immunoreactive corticotropin-releasing factor in normal subjects. Life Sci 1987;40:1651–1655. 463 Hindmarsh KW, Tan L, Sankaran K et al. Diurnal rhythms of cortisol, ACTH, and beta-endorphin levels in neonates and adults. West J Med 1989;151: 153–156. 464 Sasaki A, Shinkawa O, Yoshinaga K. Placental corticotrophin-releasing hormone may be a stimulator of maternal pituitary adrenocorticotropic hormone secretion in humans. J Clin Invest 1989;84:1997–2001. 465 Muglia LJ, Jacobson L, Weninger SC et al. Impaired diurnal adrenal rhythmicity restored by constant infusion of corticotropin-releasing hormone, in corticotropin-releasing hormone-deficient mice. Journal of Clinical Investigation 1997;99:2923–2929. 466 Muglia LJ, Jacobson L, Luedke C et al. Corticotropin-releasing hormone links pituitary adrenocorticotropin gene expression and release during adrenal insufficiency [see comments]. J Clin Invest 2000 May 2000;105:1269–1277. 467 Ellis MJ, Schmidli RS, Donald RA et al. Plasma corticotrophin-releasing factor and vasopressin responses to hypoglycaemia in normal man. Clin Endocrinol (Oxf ) 1990;32:93–100. 468 Decherney GS, DeBold CR, Jackson RV et al. Effect of ovine corticotropinreleasing hormone administered during insulin-induced hypoglycemia on plasma adrenocorticotropin and cortisol. J Clin Endocrinol Metab 1987;64: 1211–1218. 469 Adler GK, Majzoub JA. Influence of infused hypertonic saline on the response to insulin-induced hypoglycemia in man. J Clin Endocrinol Metab 1987;65: 116–121. 470 Luger A, Deuster PA, Gold PW et al. Hormonal responses to the stress of exercise. Adv Exp Med Biol 1988;245:273–280. 471 Deuster PA, Chrousos GP, Luger A et al. Hormonal and metabolic responses of untrained, moderately trained, and highly trained men to three exercise intensities. Metabolism 1989;38:141–148. 472 Janal MN, Colt EW, Clark WC et al. Pain sensitivity, mood and plasma endocrine levels in man following long-distance running: effects of naloxone. Pain 1984;19:13–25. 473 Pitts AF, Preston MA, II, Jaeckle RS et al. Simulated acute hemorrhage through lower body negative pressure as an activator of the hypothalamic–pituitary–adrenal axis. Horm Metab Res 1990;22:436–443. 474 Udelsman R, Chrousos GP. Hormonal responses to surgical stress. Adv Exp Med Biol 1988;245:265–272. 475 Udelsman R, Norton JA, Jelenich SE et al. Responses of the hypothalamic– pituitary–adrenal and renin-angiotensin axes and the sympathetic system during controlled surgical and anesthetic stress. J Clin Endocrinol Metab 1987; 64:986–994. 476 Udelsman R, Ramp J, Gallucci WT et al. Adaptation during surgical stress. A reevaluation of the role of glucocorticoids. J Clin Invest 1986;77: 1377–1381.

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Hypothalamic–Pituitary Function

477 Oltras CM, Mora F, Vives F. Beta-endorphin and ACTH in plasma: effects of physical and psychological stress. Life Sci 1987;40:1683–1686. 478 Voigt K, Ziegler M, Grunert Fuchs M et al. Hormonal responses to exhausting physical exercise: the role of predictability and controllability of the situation. Psychoneuroendocrinology 1990;15:173–184. 479 Arduini D, Rizzo G, Parlati E et al. Modifications of ultradian and circadian rhythms of fetal heart rate after fetal-maternal adrenal gland suppression: a double blind study. Prenat Diagn 1986;6:409–417. 480 Price DA, Close GC, Fielding BA. Age of appearance of circadian rhythm in salivary cortisol values in infancy. Arch Dis Child 1983;58:454–456. 481 Onishi S, Miyazawa G, Nishimura Y et al. Postnatal development of circadian rhythm in serum cortisol levels in children. Pediatrics 1983;72:399–404. 482 Attanasio A, Rosskamp R, Bernasconi S et al. Plasma adrenocorticotropin, cortisol, and dehydroepiandrosterone response to corticotropin-releasing factor in normal children during pubertal development. Pediatr Res 1987;22:41–44. 483 Ghizzoni L, Virdis R, Ziveri M et al. Adrenal steroid, cortisol, adrenocorticotropin, and beta-endorphin responses to human corticotropinreleasing hormone stimulation test in normal children and children with premature pubarche. J Clin Endocrinol Metab 1989;69:875–880. 484 McEwen BS. The neurobiology of stress: from serendipity to clinical relevance. [Review] [157 refs]. Brain Res 2000 Dec 15 2001;886:172–189. 485 Elenkov IJ, Wilder RL, Chrousos GP et al. The sympathetic nerve—an integrative interface between two supersystems: the brain and the immune system. [Review] [445 refs]. Pharmacol Rev 2000 Dec 2001;52:595–638. 486 Blalock JE. Proopiomelanocortin and the immune-neuroendocrine connection. [Review] [68 refs]. Ann NY Acad Sci 1999;885:161–172. 487 Ottaviani E, Franchini A, Genedani S. ACTH and its role in immuneneuroendocrine functions. A comparative study. Curr Pharm Des 1999;5: 673–681. 488 Elenkov IJ, Webster EL, Torpy DJ et al. Stress, corticotropin-releasing hormone, glucocorticoids, and the immune/inflammatory response: acute and chronic effects. [Review] [59 refs]. Ann NY Acad Sci 1999;876:1–11. 489 Cavagnaro J. Molecular basis for the bidirectional modulation of the neuroendocrine and the immune systems. Year Immunol 1986;2:303–322. 490 Reichlin S. Neuroendocrine-immune interactions. N Engl J Med 1993;329: 1246–1253. 491 Eskay RL, Grino M, Chen HT. Interleukins, signal transduction, and the immune system-mediated stress response. Adv Exp Med Biol 1990;274: 331–343. 492 Kavelaars A, Ballieux RE, Heijnen CJ. The role of IL-1 in the corticotropinreleasing factor and arginine- vasopressin-induced secretion of immunoreactive beta-endorphin by human peripheral blood mononuclear cells. J Immunol 1989;142:2338–2342. 493 Westly HJ, Kleiss AJ, Kelley KW et al. Newcastle disease virus-infected splenocytes express the proopiomelanocortin gene. J Exp Med 1986;163: 1589–1594. 494 Lolait SJ, Clements JA, Markwick AJ et al. Pro-opiomelanocortin messenger ribonucleic acid and posttranslational processing of beta endorphin in spleen macrophages. J Clin Invest 1986;77:1776–1779. 495 Karalis K, Sano H, Redwine J et al. Autocrine or paracrine inflammatory actions of corticotropin-releasing hormone in vivo. Science 1991;254:421–423. 496 Muglia LJ, Jenkins NA, Gilbert DJ et al. Expression of the mouse corticotropin-releasing hormone gene in vivo and targeted inactivation in embryonic stem cells. Journal of Clinical Investigation 1994;93:2066–2072. 497 Scheithauer BW. Surgical pathology of the pituitary: the adenomas. Part II. Pathol Annu 1984;19:269–329. 498 Saeger W, Geisler F, Ludecke DK. Pituitary pathology in Cushing’s disease. Pathol Res Pract 1988;183:592–595. 499 Herman V, Fagin J, Gonsky R et al. Clonal origin of pituitary adenomas. J Clin Endocrinol Metab 1990;71:1427–1433. 500 Hori T, Nishiyama F, Anno Y et al. Differences in glycoconjugates of adrenocorticotropic hormone-secretory granules between nonadenomatous pituitary cells and adenoma cells as detected by double labeling. Neurosurgery 1988;23:52–57. 501 Challa VR, Marshall RB, Hopkins MB et al. Pathobiologic study of pituitary tumors: report of 62 cases with a review of the recent literature. Hum Pathol 1985;16:873–884.

502 Apletalina V, Muller L, Lindberg I. Mutations in the catalytic domain of prohormone convertase 2 result in decreased binding to 7B2 and loss of inhibition with 7B2 C-terminal peptide. J Biol Chem 2000;275:14667–14677. 503 Schteingart DE, Chandler WF, Lloyd RV et al. Cushing’s syndrome caused by an ectopic pituitary adenoma. Neurosurgery 1987;21:223–227. 504 Axiotis CA, Lippes HA, Merino MJ et al. Corticotroph cell pituitary adenoma within an ovarian teratoma: A new cause of Cushing’s syndrome. Am J Surg Pathol 1987;11:218–224. 505 Neilson K, de Chadarevian JP. Ectopic anterior pituitary corticotropic tumour in a six-year-old boy. Histological, ultrastructural and immunocytochemical study. Virchows Arch [A] 1987;411:267–273. 506 Karl M, Lamberts SW, Koper JW et al. Cushing’s disease preceded by generalized glucocorticoid resistance: clinical consequences of a novel, dominant-negative glucocorticoid receptor mutation. Proc Assoc Am Physicians 1996;108:296–307. 507 Karl M, Von Wichert G, Kempter E et al. Nelson’s syndrome associated with a somatic frame shift mutation in the glucocorticoid receptor gene. J Clin Endocrinol Metab 1996;81:124–129. 508 Gabrilove JL, Anderson PJ, Halmi NS. Pituitary pro-opiomelanocortin-cell carcinoma occurring in conjunction with a glioblastoma in a patient with Cushing’s disease and subsequent Nelson’s syndrome. Clin Endocrinol (Oxf ) 1986;25:117–126. 509 Nawata H, Higuchi K, Ikuyama S et al. Corticotropin-releasing hormone- and adrenocorticotropin-producing pituitary carcinoma with metastases to the liver and lung in a patient with Cushing’s disease. J Clin Endocrinol Metab 1990;71:1068–1073. 510 Gould VE, Lee I, Warren WH. Immunohistochemical evaluation of neuroendocrine cells and neoplasms of the lung. Pathol Res Pract 1988;183:200–213. 511 Kasurinen J, Syrjanen KJ. Peptide hormone immunoreactivity and prognosis in small-cell carcinoma of the lung. Respiration 1986;49:61–67. 512 Sieber SC, Gelfman NA, Dandurand R et al. Ectopic ACTH and adrenal myelolipoma. Conn Med 1989;53:7–10. 513 Herbst WM, Kummer W, Hofmann W et al. Carcinoid tumors of the thymus. An immunohistochemical study. Cancer 1987;60:2465–2470. 514 Arioglu E, Doppman J, Gomes M et al. Cushing’s syndrome caused by corticotropin secretion by pulmonary tumorlets. N Engl J Med 1998;339:883–886. 515 Raux Demay MC, Proeschel MF, de Keyzer Y et al. Characterization of human corticotrophin-releasing hormone and pro-opiomelanocortin-related peptides in a thymic carcinoid tumour responsible for Cushing’s syndrome. Clin Endocrinol (Oxf ) 1988;29:649–657. 516 Suda T, Kondo M, Totani R et al. Ectopic adrenocorticotropin syndrome caused by lung cancer that responded to corticotropin-releasing hormone. J Clin Endocrinol Metab 1986;63:1047–1051. 517 Sakurai H, Yoshiike Y, Isahaya S et al. A case of ACTH-producing pheochromocytoma. Am J Med Sci 1987;294:258–261. 518 Mendonca BB, Arnhold IJ, Nicolau W et al. Cushing’s syndrome due to ectopic ACTH secretion by bilateral pheochromocytomas in multiple endocrine neoplasia type 2A [letter]. N Engl J Med 1988;319:1610–1611. 519 Southgate HJ, Archbold GP, el Sayed ME et al. Ectopic release of GHRH and ACTH from an adenoid cystic carcinoma resulting in acromegaly and complicated by pituitary infarction. Postgrad Med J 1988;64: 145–148. 520 Watanobe H, Yoshioka M, Takebe K. Ectopic ACTH syndrome due to Grawitz tumor. Horm Metab Res 1988;20:453–456. 521 Werner S, Jacobsson B, Bostrom L et al. Cushing’s syndrome due to an ACTH-producing neuroendocrine tumour in the nasal roof. Acta Med Scand 1985;217:235–240. 522 Long SI. ACTH-producing apudoma metastatic to the liver. J Natl Med Assoc 1987;79:122–123. 523 de Keyzer Y, Bertagna X, Luton JP et al. Variable modes of proopiomelanocortin gene transcription in human tumors. Mol Endocrinol 1989;3: 215–223. 524 Melmed S, Yamashita S, Kovacs K et al. Cushing’s syndrome due to ectopic proopiomelanocortin gene expression by islet cell carcinoma of the pancreas. Cancer 1987;59:772–778.

C h a p t e r

3 Growth Hormone Vivien S. Herman-Bonert Shlomo Melmed

GROWTH HORMONE (GH) GENE STRUCTURE

Somatotroph Development and Differentiation

The human GH genomic locus spans approximately 66 kb and contains a cluster of five highly conserved genes [1] located on the long arm of human chromosome 17 at bands q22–24 [2]. The 5¢ to 3¢ arrangement of these genes is hGHN, hCS-L, hCS-A, hGH-V, and hCS-B [1], all of which have the same basic structure consisting of five exons separated by four introns [1]. The hGH-N gene is transcribed only in somatotrophs of the anterior pituitary while the hCS-A and hCS-B genes are expressed in placental trophoblasts [3]. hGH-N codes for a 22-kDa protein consisting of 191 amino acids. Approximately 10% of pituitary GH is presented as a 20-kDa variant lacking amino acid residues 32–46 [4,5], and probably arising as a result of an alternate splicing mechanism [6]. hGH-V is expressed by the syncytiotrophoblast of the placenta during the second and third trimesters of gestation [7,8], hGH-V messenger RNA (mRNA) encodes 22kDa protein secreted form which can be detected in the maternal circulation from midpregnancy [9] and a minor form hGH-V2 which is predicted to be a 26-kDa protein product [10]. The role of hGH-V is unknown, however, the rise in hGH-V concentrations in maternal serum correlates with a fall in hGH-N concentrations, suggesting the possibility of a feedback loop on the maternal hypothalamic pituitary axis [9]. Postpartum, GH-V levels drop rapidly and are undetectable in the circulation after one hour [9]. hGH-V has a greater binding affinity than hGH-N for somatogen vs lactogen receptors [11]. hGH-V and hGH-N also differ in their ratio of somatogen to lactogen bioactivities, with hGH-V having the greater ratio [12]. In addition, hGH-V has been shown to influence carbohydrate and fat metabolism in rat adipose tissues in a manner similar to that of hGH-N [13]. The hCS-L gene is not known to yield a product [14].

GH is specifically expressed in the somatotroph cell of the anterior pituitary which develops in a time-and-space dependent manner. Expression of the a-subunit transcript in the hypophyseal placode within ectoderm of the pharynx prior to the formation of Rathke’s pouch defines the onset of pituitary organogenesis [15]. The mammalian anterior pituitary develops from Rathke’s pouch during the early stages of embryonic development [16]. Its ventral epithelium serves as the anterior pituitary anlagen while the dorsal epithelium generates the intermediate lobe of the pituitary. A process of cytodifferentiation gives rise to the different hormone-producing cells [17]. Acidophils are the progenitors for both GH-producing somatotrophs and prolactin producing lactotrophs (PRL), while basophils give rise to cells which secrete adrenocorticotrophic hormone (ACTH), thyroid-stimulating hormone (TSH), luteinizing hormone (LH), and follicle-stimulating hormone (FSH) [17]. Cells of the intermediate lobe produce pro-opiomelanocortin (POMC) gene-related peptides [17]. Experimental evidence obtained from transgenic mice studies suggests that most PRL-expressing cells arose from GH-producing cells [18]. Ablation of somatotrophs by expression of GH-diphtheria toxin and GH-thymidine kinase fusion genes inserted into the germ line of transgenic mice results in the elimination of the majority of lactotrophs; however a small percentage of lactotrophs escape destruction [18,19]. This suggests that the majority of PRLproducing cells arose from postmitotic somatotrophs. The Pit-I gene transcript and POUIFI protein are found in somatotrophs, lactotrophs and thyrotrophs [15]. This suggests an mRNA-specific translational control as part of the developmental process [15]. The actions of Pit-1 protein are complemented by other factors required to achieve the 79

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physiologic patterns of cell-specific gene activation [15]. The estrogen receptor, which is activated subsequent to the appearance of Pit-1, appears to mediate in part a progressive increase in PRL gene expression characteristic of mature lactotrophs. Inherited isolated GH deficiency and short stature may be attributed to mutations of transcription factors, the GH-1 gene, the GHRH receptor, the GH receptor or rarely IGF-related molecules.

Promoter Structure The hGH promoter region contains cis-elements which mediate both pituitary-specific and hormone-specific signaling [20,21]. DNase footprinting reveals several proteinbinding sites within 300 bp of the 5¢ flanking DNA. One one of these proteins is unique to pituitary cells, suggesting that it plays a major role in tissues-specific expression of the GH gene [20,22]. This factor, alternately named Pit-1 or GHF-1 [23,24], binds to two sites centered around sequences -80 and -120. Pit-1 is a 31- or 33-kDa protein important for GH promoter activity both in vivo and in vitro [23–25]. When transferred into nonexpressing cells, Pit-1 is capable of stimulating GH-directed gene expression in nonpituitary cells [20]. In addition, Pit-1 binds to the PRL promoter [26]. Pit-1 is a member of a family of transcription factors sharing two regions of homology [27]; a highly conserved POU-specific (POUs) domain and a more divergent homeodomain (POUHD). Initial recognition of this family resulted from the cloning of complementary DNAs (cDNAs) encoding several transactivators Pit-1 [23,24], Oct1 [28,29], Oct-2 [30,31], and unc-86 [32]. The POUs domain is unique to this family of proteins but the POUHD, which is divergent, shares sequence homology and the predicted a-helical structures with classic homeodomain-containing proteins [33]. Data from mutant Pit-1 proteins suggest that while POUHD is required and sufficient for low-affinity DNA binding, the POUs domain is necessary for highaffinity binding and accurate recognition of natural Pit-1 response elements. In addition, POU domain proteins, in contrast to homeodomain proteins, require that both the POUs and POUHD contact the DNA [27]. Transcriptional activity of Pit-1 is conferred by a serine- and threoninerich N-terminal region which is different to the transcriptional contacts of other types of activation domains [27]. A variant isoform of Pit-1, Pit-1B [34], contains a 26-amino acid insertion in the transactivation domain and, like the previously described Pit-I protein, is capable of inducing both GH and PRL promoters. PROP-1, a paired homeobox protein, is required for initial commitment of Pit-1 cell lineages [35]. PROP-1 represses Rpx expression, and missense and spliced mutations of PROP-1 leading to loss of DNA-binding or transactivation leads to pituitary failure with short stature and varying degrees of thyroid failure, hypogonadism, and ACTH deficiency [36].

POUIFI [37] mutations may also lead to pituitary failure. Patients with combined pituitary hormone deficiency have predominantly GH and PRL deficiency, with variable degrees of hypothyroidism [37]. Footprinting analysis reveals the presence of a second, ubiquitous factor binding to the distal Pit-1 site [38,39]. The protected template contains a consensus sequence for the binding of Sp1, a transcription factor that recognizes the glycine-cysteine box in several viral and eukaryotic gene promoters [40–42]. Pit-1 binds to its distal site with a lower affinity [23,24], and, in addition, Pit-1 and Sp1 bind in a mutually exclusive manner to the GH promoter [38,39]. In vivo, both binding sites contribute to rat GH promoter activation [39]. Mutation of the Sp1 binding site results in a 50% reduction of promoter activity, suggesting that activation of GH by Pit-1 and Sp1 may occur through a multistage mechanism. Sequences -266/-252 of the hGH promoter contain an 8 bp recognition site for the adenoviral major late transcription factor (MLTF), also termed upstream stimulating factor (USF) [43]. A USF-like protein also binds to the upstream hGH promoter [38,43,44]. This factor contributes to one of the protected sites in the third footprint of the hGH gene promoter. USF is a basal transcription factor initially identified in Hela cell extracts [45,46]. Known targets of USF include adenovirus-2 [45,46], fibrinogen-g and the metallothionein-1 genes [47,48]. The distal part of this protected region of the hGH promoter, -290/-272, binds an insulin-inducible nuclear factor [44] that may represent an NF1 family protein [49]. A locus control region (LCR) of the hGH gene located 5¢ to the transcription start site determines somatotroph and latotroph GH expression. This genetic control also involves appropriate regulation of a chromatin domain in these pituitary cell types [50].

Hormonal Regulation of the hGH Gene Synthesis and release of GH is under control of a variety of hormonal agents, including insulin growth factor-I (IGF-I), GH-releasing hormone (GHRH), Ghrelin, somatostatin, thyroid hormone, glucocorticoids, and insulin. IGF-I inhibits GH mRNA expression as well as GH secretion in dispersed human adenoma cell cultures [51]. This effect is blocked by the monoclonal IGF-I receptor antibody a-IR3 [51]. IGF-I also attenuates basal and stimulated GH gene expression when 500 bp of the hGH gene promoter was transfected into human choriocarcinoma cells ( JEG3) or HeLa cells [52]. GHRH stimulates the release of hGH, and cyclic adenosine monophosphate (cAMP) is thought to be the second messenger for this effect. Analysis of the hGH promoter for a cAMP-responsive element (CRE) fails to reveal a consensus sequence. Two groups have reported the presence of novel CREs within -83 and -150 bp of the 5¢ flanking DNA, respectively [21,53]. CREB binding protein (CBP) is also a co-factor for Pit-1 dependent human GH

Chapter 3

activation. CBP is phosphorylateral by protein kinase A, independently of CREB [54]. The role of thyroid hormone in hGH synthesis remains unclear. Release of hGH in response to provocative stimuli is usually reduced in hypothyroid patients; however, basal GH levels and responses may be normal [55,56]. In numerous hGH gene transfer studies, absence of a T3 response has been documented [57,58]. Glucocorticoid responsiveness of the hGH gene has been demonstrated with the strongest receptor binding site being located within the first intron of the hGH gene; however, a 5¢ GRE is also present and capable of weak induction [59–61]. Insulin suppresses hGH secretion by acromegaly tumor cells in culture [62], and also attenuates basal and stimulated transient expression of a transfected hGH gene [63]. Somatostatin is synthesized in the medial preoptic area of the hypothalamus and inhibits secretion of GH from human adenoma cell cultures [64]. Thus, numerous ubiquitous factors bind to the GH promoter. In the absence of Pit-1 the promoter region is inactive, and binding of Pit-1 to the transcriptional machinery facilitates the interaction between this factor and other ubiquitous activators already bound to the promoter or subsequently activated to enhance GH transcription. This model

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81

of cooperative interaction provides an explanation for tissuespecific expression of the hGH gene by a single cell typespecific activator. CONTROL OF GH SECRETION The GH molecule, a single-chain polypeptide hormone consisting of 191 amino acids (Fig. 3.1), is synthesized, stored, and secreted by somatotroph cells. The crystal structure of human GH and GH-binding protein reveals four alpha helixes, and two identical GHBP molecules occurring within a single GH molecule [65]. Somatotrophs are located predominantly in the lateral wings of the anterior pituitary gland. Somatotrophs comprise 35% to 45% of pituitary cells and the gland contains a total of 5–15 mg of GH [66]. Circulating GH molecules are a heterogenous group of peptides [67] which comprises at least three monomeric forms and several oligomers. The monomeric moieties include a 22- and 20-kDa form, acetylated 22K, and two desamido GH’s. The 22-kDa peptide is the major physiologic GH component. The 20-kDa GH has a slower metabolic clearance than the 22-kDa form [68], which accounts for the plasma 20 : 22 ratio being higher than in the pituitary gland.

FIGURE 3.1. Amino acid structure of human growth hormone (GH). GH is a 191amino acid single-chain 21.5 kDa polypeptide with two intramolecular disulfide bonds. Fifteen percent of GH is deleted from amino acid (32–46) and is secreted as a 20 kDa protein.

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The 22-kDa peptide retains growth-promoting activity, but lacks diabetogenic effects, which are more pronounced with the 20-kDa form. The relative proportions of circulating GH molecular forms in children of various ages are quantitatively and qualitatively identical in adults. Monomeric GH forms found in the plasma of acromegalic patients are also qualitatively similar to those found in normal plasma [69].

Ontogeny of GH Levels Circulating GH is first detectable in fetal serum at the end of the first trimester, peaks at a concentration of 100 to 150 ng/ml at 20 weeks of gestation, and subsequently falls to 30 ng/ml at birth. GH levels continue to fall during infancy. During childhood, levels are similar to those in adulthood, until puberty, when circulating levels are elevated. GH levels decline after adolescent growth and remain stable until mid-adulthood, when they decline progressively through old age [70]. Decreased sensitivity of the pituitary to GHRH action with aging may contribute to these GH fluctuations with advancing age [71].

Physiologic Factors Involved in GH Secretion GH secretion is pulsatile, the anterior pituitary gland secreting bursts of GH, with almost undetectable basal levels occurring between these peaks [72]. The number of GH pulses detected depends on the frequency of blood sampling. When sampling frequency is reduced to three or fewer samples per hour, less than 0.4 GH pulses/hr are detected, and this rate increases as sampling is intensified [72]. Integrated GH levels are higher in women than in men [73], and are also enhanced in postmenopausal women following estrogen replacement [74]. In children and young adults, maximum GH secretion occurs within an hour of the onset of deep sleep (stage III or IV), with subsequent smaller peaks appearing during later sleep [75]. Effects of Sleep

A major GH secretory pulse occurs shortly after the onset of sleep, associated with the first episode of slow wave sleep [76]. Sleep stimulates GH secretion and 60% to 70% of daily GH secretion occurs during early sleep, in association with slow wave sleep [77] Rapid eye movement (REM) sleep is reduced by approximately 50% after age 50 years with significant sleep fragmentation [78]. The decline in slow wave sleep from early adulthood to midlife is paralleled by a major decline in GH secretion. This suggests that the age-related alterations in the somatotropic axis may partially reflect decreased sleep quality. Although increased PRL and ACTH concentrations occur later during sleep, their secretion is not as tightly linked to sleep patterns as is GH secretion. “Jet lag” transiently increases the height of GH peaks during the day and night, resulting in a transient increase of 24-hour

GH secretion. Jet lag also shifts the major GH secretory spike from early to late sleep [79]. Exercise

Exercise increases GH secretion probably mediated by a cholinergic mechanism [80]. Stress

GH release is stimulated by physical stress, including trauma with hypovolemic shock [81] and sepsis [82]. However, chronic debilitating diseases, including cancer, are not associated with increased GH levels [83]. Increased GHRH release, mediated by adrenergic pathways, is thought to mediate stress-induced GH secretion [84]. Emotional deprivation is associated with suppressed GH secretion [85], and subnormal GH responses to provocative stimuli have been described in endogenous depression [86]. Nutritional and Metabolic Effects

Nutritional and metabolic factors profoundly influence GH secretion. Chronic malnutrition [87] and voluntary 5-day fasting [88] are associated with elevated GH levels. Both pulse frequency and amplitude of GH secretory peaks increase with fasting [88] (Table 3.1). Obesity decreases basal and stimulated GH secretion. Insulin-induced hypoglycemia stimulates GH release 30 to 45 minutes after the glucose trough, whereas acute hyperglycemia inhibits GH secretion [89]. Diabetic patients with chronic hyperglycemia, however, do not have suppressed GH levels and in fact many poorly controlled diabetic patients have increased basal [90] and exercise-induced GH levels [91]. Central nervous system glucoreceptors appear to sense fluctuations, rather than absolute glucose levels. However, glucose homeostasis is not the major determinant of GH secretion, this being overridden by effects of sleep, exercise, stress, and by random GH bursts. A high-protein meal, and single amino acids (including arginine and leucine) administered intravenously stimulate GH secretion. Arginine may suppress endogenous somatostatin secretion and thereby stimulate GH secretion [92]. Decreased serum free fatty acid (FFA) levels cause acute GH release [93] and increased serum FFA blunt the effects of various stimuli, including arginine infusion, [94] sleep, Ldopa, and exercise [95–98] on GHRH-stimulated GH release [98].

Table 3.1. Deconvolution analysis of growth hormone (GH) secretion in adult males. From Thorner et al. [88a]

24 hr secretion Secretory bursts GH/burst

Normal

Fasting

Obesity

Middle-age

540 ± 44 12 ± 1 45 ± 4

2171 ± 333 32 ± 2 64 ± 9

77 ± 20 3 ± 0.5 24 ± 5

196 ± 65 10 ± 1 20 ± 6

Chapter 3

Growth Hormone

83

FIGURE 3.2. Schematic representation of the actions of leptin. Leptin acts either directly or by activating specific centers in the central nervous system to decrease food intake, increase energy expenditure, influence glucose and fat metabolism, and alter neuroendocrine function. From Mantzoros [100a].

FIGURE 3.3. Schematic representation of feedback loops involving leptin. Leptin, an adipocyte-derived hormone, circulates in the serum either in free form or bound to leptinbinding proteins, activates specific receptors in the hypothalamus, and alters expression of several neuropeptides; these in turn decrease appetite, increase energy expenditure by altering sympathetic and parasympathetic tone, and alter neuroendocrine function. Increasing leptin levels activate the thyroid, growth hormone, and gonadal axes and suppress the pituitary–adrenal axis. Leptin, acting directly or indirectly (by altering the levels of other hormones and neuropeptides), also influences hemopoiesis and immune function and improves glucose and fat metabolism. From Mantzoros [100a].

Leptin

Leptin, a 167 amino acid cytokine, is the product of the ob gene, expressed in white adipose tissue, stomach, placenta and possible mammary gland [99]. Leptin is thought to play a key role in the regulation of body fat mass [100]. Regulating food intake and energy expenditure (Fig. 3.2). GH has several metabolic effects, including lipolytic and anabolic effects. In turn, alterations in nutritional status, such as obesity and food deprivation, influence GH secretion. In obese subjects, GH secretion is markedly impaired. The mechanism by which metabolic status regulates GH secretion is poorly understood. As leptin receptors are present on

the hypothalamus, and leptin plays a role in hypothalamic–pituitary function in fasted animals [101], leptin may act as a metabolic signal to regulate GH secretion (Fig. 3.3). Leptin antiserum administered to normal fed rats, decreased plasma GH levels [102], implying that physiologic leptin levels may facilitate normal spontaneous GH secretion. Furthermore leptin administered to fasted rats reversed the inhibitory effect of food deprivation on in vivo GH secretion [102]. Increased SRIF release may be a factor in the suppressed GH secretion observed in fasted rats [103], possibly mediated by neuropeptide Y, as hypothalamic neuropeptide Y

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release is increased in fasted rats [104]. Leptin receptors are present in neuropeptide Y producing hypothalamic neurons [105], which synapse with somatostatin neurones [104]. Furthermore both antisera to neuropeptide Y and somatostatin reverse starvation-induced GH release [103,104]. Further studies are needed to elucidate the effects of leptin on neuropeptide Y, and somatostatin and its role as a metabolic signal in the regulation of GH secretion. In GH deficient hypopituitary adults, leptin concentrations are more elevated than would be expected from the degree of obesity [106].

Cholinergic and Serotonergic Neurotransmitters

Cholinergic and serotoninergic neurons have been implicated in the etiology of sleep-induced GH secretion [119]. Opiods

Endorphins and enkephalins stimulate GH secretion in animals [120], probably via interaction with other neurotransmitters in both the pituitary and hypothalamus. Opioids increase rat GHRH concentrations [121], and endogenous opiates probably increase GH release during severe physical stress and extreme exercise [122]. Gastrointestinal Neuropeptides

Pharmacologic Control of GH Secretion The central neurogenic control of GH is complex. Neuropeptides, neurotransmitters and opiates impinge on the hypothalamus and modulate GHRH and somatostatin (SRIF) release. The net effect of these complex influences determines the final secretory pattern of GH.

Several gastrointestinal neuropeptides stimulate GH secretion in animal models, including substance P, neurotensin, vasoactive intestinal polypeptide, peptide histidine isoleucine amide (PHI), motilin, galanin, cholecystokinin, and glucagon [123]. HYPOTHALAMIC HORMONES

Dopaminergic Neurotransmitters

Dopamine (DA) is a precursor of epinephrine and norepinephrine. Apomorphine, a central dopamine receptor agonist, stimulates GH secretion [107]. GH-deficient children have been shown to increase their growth velocity after 6 months of levodopa treatment [108]. Sixty to 90 minutes after oral l-dopa administration, adults increase their serum GH levels from 0 to 5–20 ng/ml [109]. Phentolamine, an a-adrenergic blocking agent, inhibits l-dopa-induced GH release, indicating that l-dopa may, in addition, require conversion to epinephrine or norepinephrine to exert its effect on GH secretion [110]. Catecholaminergic Neurotransmitters

Norepinephrine increases GH secretion via a-adrenergic pathways and inhibits GH release via b-adrenergic pathways [111]. Insulin-induced hypoglycemia increases GH secretion via an a2-adrenergic pathway [112], whereas clonidine acts on a1-adrenergic receptors to increase GH secretion [113]. Arginine administration, exercise, l-dopa, and antidiuretic hormone (ADH) facilitate GH secretion by a-adrenergic effects [114]. Animal studies using pharmacologic agents and passive immunization show that a-adrenergic blockade simultaneously inhibits GHRH release while stimulating SRIF release, resulting in suppressed GH secretion [115]. b2-adrenergic receptor stimulation decreases basal GH levels [116]. b-adrenergic blockade increases GHRH-induced GH release, possibly due to a b-adrenergic effect at the pituitary level or via decreased hypothalamic somatostatin release [117]. b-blockade also enhances GH release elicited by insulin-induced hypoglycemia, ADH, glucagon, and L-dopa [114]. Epinephrine may regulate GH release by decreasing somatostatin release [118], and metabolites of g-aminobutyric acid (GABA) may alter GH secretion by enhancing catecholaminergic release [110].

Thyrotropin-releasing hormone (TRH) does not stimulate GH secretion in normal subjects [124]; it does, however, induce GH secretion in about 70% of patients with acromegaly [125]. TRH also stimulates in vitro GH secretion in cultured pituitary tumor cells [126]. The discordant GH response to TRH has also been demonstrated in patients with liver disease [127], renal disease, ectopic GHRH-releasing carcinoid tumors [128], anorexia nervosa [129], and depression [130]. Administration of intravenous corticotropin-releasing hormone (CRH) increases GH secretion in a small percentage of patients with acromegaly [131] and in patients with chronic depression [132]. Gonadotropin-releasing hormone (GnRH) stimulates GH secretion in one-third of patients with acromegaly, but the mechanism is unclear [133].

Hormones Facilitating GH Secretion Physiologic glucocorticoid concentrations increase GH secretion in human pituitary cultures [134]. In acromegalic patients, dexamethasone suppresses GH secretion in vivo [135]. Supraphysiologic serum glucocorticoid concentrations retard growth in humans [136]. Cushing’s disease is also associated with growth retardation, decreased serum GH [137], and decreased pituitary GH content in the tissue surrounding the adenoma [138]. A single dose of dexamethasone administered to normal subjects suppresses GHRH-induced GH release within 12 hours. Glucocorticoids administered to normal subjects produce a dose dependent inhibition of GHRH-stimulated GH secretion, identical to that seen in Cushing’s syndrome [139,140]. Glucocorticoids have a greater inhibitory action on GHRHinduced GH secretion than clonidine or arginine [139]. In chronic hypocortisolism, GHRH-induced GH secretion is reduced [141]. To elucidate the mechanism of the blocking

Chapter 3

action of glucocorticoids on GH release, increased somatostatin secretion seems unlikely, as pyridostigmine (which blocks hypothalamic somatostatin secretion) does not increase suppressed GH levels in Cushing’s syndrome [142]; neither can IGF-I be incriminated, as IGF-I levels are normal in Cushing’s syndrome [143]. The above data exclude decreased endogenous GHRH release as the mechanism. When the glucocorticoid effect is measured acutely, i.e., at 3 hours, stimulated GH levels are measured, which remain elevated for 2 hours [144]. Acute glucocorticoidinduced GH secretion occurs with several different corticosteroid preparations including hydrocortisone, deflazacort, and dexamethasone. The short-term stimulatory effect of glucocorticoids on GH secretion is one of the most potent and also the most delayed stimulus to GH secretion, as it takes 3 hours to occur [144]. Thus glucocorticoids have short-term stimulatory effects on GH secretion and delayed inhibitory effects. Specific glucocorticoid responsive elements have been identified on the GH gene. Currently, the mechanism of action of the dual stimulation and inhibition of glucocorticoid on GH secretion has not been elucidated. Thyroid hormone increases basal and stimulated rat GH synthesis, acting directly at the level of gene expression [145]. This effect appears to be synergistic with glucocorticoids. T3 binds to nuclear receptors in rat pituitary tumor cells [146] and stimulates GH gene transcription [147]. In vitro data confirm that the hGH promoter is not thyroid hormone-responsive, while T3 strongly induces the rat GH promoter [21,148]. In hyperthyroid patients, GH secretion rate is decreased, but normalizes when patients are rendered euthyroid [149], suggesting that thyroid hormone suppresses GH secretion from the normal human somatotroph in vivo. GHRH

Hypothalamic GHRH was characterized initially by two different groups from ectopic pancreatic GHRH-secreting tumors causing acromegaly [128,150]. Analysis of one tumor revealed a 44-amino acid GHRH residue [151]; the other contained 37-, 40-, and 44-amino acid forms [152]. GHRH (1–40) and GHRH (1–44) are both found in extracts derived from the human hypothalamus. GHRH is secreted

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85

from neurons in the hypothalamic arcuate nucleus and premammillary area, with axons that project to the median eminence [153]. The hGHRH gene encodes a 108-amino acid prepro-hormone for GHRH-44 [154,155], which has a free amino terminal and amidated carboxy terminal residue. The amino terminal tyrosine appears to bestow biologic activity on the GHRH molecule (Fig. 3.4). There is considerable structural homology between GHRH and several gut peptides. The highest is between GHRH and PHI, which have 12 amino acids in common in equivalent positions [156]. Varying degrees of homology exist between GHRH and VIP, glucagon, secretin, and GIP [157]. All of these peptides stimulate GH secretion in various physiologic systems, but with lower potency than GHRH. GHRH binds to specific sites on the somatotroph membrane, resulting in increased intracellular 3¢, 5¢cAMP [158]. The GHRH receptor gene has recently been cloned and sequenced, encoding a 47-kDa protein of 423 amino acids [159]. GHRH has a selective action on GH release; it does not release other anterior pituitary or gut hormones. GHRH increases GH synthesis as well as secretion, and stimulates transcription of GH mRNA [64,160]. GHRH stimulates GH release from both stored and newly synthesized intracellular GH pools, with a greater effect on stored pools [161]. There are a number of heterogeneous GH pools, varying according to time of GH synthesis [162–164], molecular size [163], and response to provocative stimuli [163]. Somatostatin suppresses both basal and GHRHstimulated GH release, but does not affect GH biosynthesis [165]. It is thus apparent that GHRH stimulates both release and synthesis of GH independently of any effects of somatostatin. GHRH administered to normal adults elicits a prompt increase in serum GH levels, with higher levels occurring in female subjects [166]. Most acromegalic patients retain an intact GH response to administered GHRH [167]. SRIF

SRIF, a cyclic tetradecapeptide, is recognized as a heterogeneous group of molecules including quantitatively predominant, but less bioactive SRIF-14, more bioactive SRIF-28,

FIGURE 3.4. Amino acid structure of growth hormone-releasing hormone (GHRH) (1–44). GHRH (1–44) and GHRH (1–40) have similar GH-releasing potency in vivo. The tyrosine at the amino terminus confers biologic activity. Fragments as short as GHRH (1–29) exhibit full biologic activity. The carboxy terminal fragment (28–44) has no biologic activity.

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FIGURE 3.5. Amino acid structure of native somatostatin (SRIF). SRIF, a tetradecapeptide (14-amino acid peptide) designated SRIF-14, SRIF-28, which is extended at the amino terminal, and several larger forms range in size from 11.5 to 15.7 kDa.

sst

PHOSPHATASE

ADENYLCYCLASE K/Ca++ CHANNELS cAMP

Ca++

Calcineurin

Ser/Thr Phosphatase

– Exocytosis

INHIBITS SECRETION

FIGURE 3.6. SRIF receptor signaling pathway: somatostatin receptors mediate their responses via several cellular effectors such as adenylyl cyclase potassium and calcium channels and phosphatases. Adapted from Yogesh et al. [176a].

SRIF block the GH secretory response to insulin-induced hypoglycemia, exercise, arginine, morphine, levodopa, and sleep related GH release. Somatostatin exerts its biologic effects through specific membrane-bound high affinity receptors. At least five somatostatin receptor (SSTR) subtypes have been cloned [176], termed SSTRs1-5. Somatostatin receptors are coupled to guanine nucleotide protein (G), and have seven transmembrane domains. There is 42% to 60% amino acid homology among the five somatostatin receptor subtypes [177]. Somatostatin receptors mediate their responses via several cellular effectors such as adenylyl cyclase, protein phosphatases, Na+-H+ exchanger, cyclic GMP-dependent protein kinases, phospholipase C, potassium and calcium channels [176] (Fig. 3.6). The human pituitary gland expresses SSTR1, 2 and 5 [178,179], whereas human pituitary adenomas contain SSTR1, 2, 3 and [178–181]. Somatostatin analogs, octreotide and lanreotide, used to control GH hypersecretion in acromegaly, bind with high affinity to SSTR 2 and less efficiently to SSTR5 [182]. SSTR1-4 have highter affinity for SRIF-14, whereas SSTR5 bind more potently to SRIF-28 [176] (Table 3.2). GHRH and SRIF Interaction in Regulating GH Secretion

and still larger forms which range in molecular mass from 11.5 to 15.7-kDa [168] (Fig. 3.5). The SRIF precursor is a 116-amino acid prohormone consisting of a 24-amino acid signal peptide, a 64-amino acid connecting region, followed by SRIF-28 [169] which incorporates SRIF-14. Prosomatostatin is synthesized in neuronal perikarya of the anterior hypothalamic periventricular nuclei, and transported by axonal flow to the nerve terminals ending near the hypophyseal portal vessels. SRIF has also been isolated from pancreatic islets, gastrointestinal, neural and epithelial cells, and extrahypothalamic central nervous system neurons [170]. SRIF has a short plasma half-life of 2 to 3 minutes [168]. SRIF inhibits GH secretion, TSH release, TRH stimulation of TSH but not PRL [171], and pancreatic secretion of insulin and glucagon [172]. Dopamine [173], substance P, and neurotensin [174] stimulate SRIF release from the hypothalamus, while glucose inhibits SRIF secretion in cultured rat hypothalamus [174]. SRIF-28 binds to normal rat pituitary receptors with a threefold greater affinity than SRIF-14 [175]. Both SRIF14 and SRIF-28 block the effect of GHRH on GH release in vitro and in vivo in many different species. Infusions of

SRIF and GHRH secreted in independent waves from the hypothalamus interact to generate pulsatile GH release. SRIF inhibits GH secretion, while GHRH stimulates GH synthesis and secretion. GHRH stimulates pituitary GH synthesis and secretion, and also induces transcription of the GH gene [160]. SRIF suppresses GH secretion without altering GH mRNA levels [64]. GH secretion is further regulated by its target growth factor, IGF-I, which participates in a hypothalamic–pituitary–peripheral regulatory feedback system [183]. GH stimulates the liver and other peripheral tissues to produce IGF-I, which exerts a feedback effect on the hypothalamus and pituitary. IGF-I stimulates hypothalamic SRIF release [172] and inhibits pituitary GH gene transcription and GH secretion [51,184]. In vitro, IGF-I directly attenuates hGH gene expression [51]. Specific antibodies against GHRH and SRIF have been used to dissect the respective contributions of SRIF and GHRH in the generation of GH pulsatility in rats. Anti-SRIF antibody administration results in elevated baseline GH levels, with intact intervening GH pulses [185]. These studies imply that hypothalamic SRIF secretion generates GH troughs. AntiGHRH antibodies eliminate spontaneous GH surges. In humans, GH pulsatility persists when GHRH is tonically

Chapter 3 Table 3.2. receptors

Growth Hormone

87

Properties of the five cloned subtypes of human somatostatin

Properties

Somatostatin receptor Subtype 1

Subtype 2

Subtype 3

Subtype 4

Chromosomal location

14

17

22

20

16

G protein coupling

Yes

Yes

Yes

Yes

Yes

Effector system Adenylate cyclase activity Tryosine Phosphatase activity

Reduced Increased

Reduced Increased

Reduced ?

Reduced ?

Reduced ?

IC50 (nM) Somatostatin-14 Somatostatin-28 Octreotide Vapreotide (RC-160) Lanreotide (BIM-23014)

1.1 2.2 >1000 >1000 >1000

1.3 4.1 2.1 5.4 1.8

1.6 6.1 4.4–35 30.9 43

0.5 1.1 >1000 45 66

0.9 0.07 5.6 0.7 0.6

Brain, lungs, jejunum, kidneys, liver and pancreas

Brain and kidneys

Brain and pancreas

Brain and lungs

Brain, heart, adrenal glands placenta, pituitary, small intestine and skeletal muscle

Distribution in normal human tissue

Subtype 5

Adapted from Patel YC and Srikant CB [182].

elevated due to ectopic GHRH production by a tumor or during GHRH infusion [127,186], suggesting that hypothalamic SRIF is mainly responsible for GH pulsatility. The rat hypothalamus releases GHRH and SRIF 180° out of phase every 3 to 4 hours, resulting in pulsatile GH levels [185]. GHRH and SRIF also act synergistically, in that preexposure to SRIF has been shown to enhance somatotroph sensitivity to GHRH stimulation [187]. Hence, during a normal GH trough period, the high SRIF level probably primes the rat somatotroph to respond maximally to the subsequent GHRH pulse, thus optimizing GH release. In addition to exerting opposing and cooperative interactions on the pituitary somatotroph, SRIF exerts a central inhibitory effect on GHRH release via direct synaptic connections between SRIF-containing axons and GHRHcontaining perikarya in the rat hypothalamic arcuate nucleus [188]. GH Autoregulation

Chronic GHRH stimulation, either by continuous infusion or repeated bolus administration, eventually results in decreased GH release from cultured rat anterior pituitary cells [189] and in vivo [190] due to somatotroph desensitization. Loss of GH sensitivity to administered GHRH does not occur in acromegalic patients in vivo [191] or in somatotroph adenomas in vitro [192], possibly reflecting larger intracellular pools of GH or abnormal signaling. One possible mechanism of pituitary desensitization is depletion of a GHRH-sensitive pool of GH. GHRH pretreatment in

vitro also leads to a 50% decrease in somatotroph GHRH binding sites [193]. Feedback loops exist between GH and IGF-I and the release of SRIF and GHRH (Fig. 3.7). GH stimulates hypothalamic SRIF release in vitro [194], and in vivo, GH administration decreases GH responses to GHRH [195], most likely by increasing hypothalamic SRIF release [196]. GHRH and SRIF also autoregulate their own secretion. GHRH inhibits its own secretion but stimulates SRIF secretion in vitro [197], while SRIF inhibits its own secretion in vitro [198]. Growth Hormone Secretagogues (GHS) and Ghrelin

Small synthetic molecules termed growth hormone secretagogues (GHS) [199] stimulate and amplify pulsatile pituitary GH release [200], via a separate pathway distinct from GHRH/SRIF. A strategy of expression cloning was used to identify the GHS receptor, which is a heterotrimeric GTP-binding protein (G-protein)-coupled molecule [201]. This classic Gprotein coupled receptor has seven alpha helical membrane spanning domains and three intracellular and extracellular loops. The GHS receptor is most strongly expressed in the anterior pituitary and both the hypothalamic and nonhypothalamic regions of the brain. The ligand of the GHS receptor is a 28 amino acid peptide, Ghrelin isolated from stomach, with 3 serine residues and is n-octanoylated [202]. It releases GH both in vivo and in vitro and 0-n octanoylation appears to be essential for GH releasing activity. The

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FIGURE 3.7. Growth hormone (GH), insulin-like growth factor-I (IGF-I) Growth Hormone secretagogues GHS (GHRH and Ghrelin), somatostatin (SRIF) feedback loop in the control of somatic growth. A stimulatory effect of GHRH and Ghrelin and inhibitory effect of SRIF interact to result in GH secretion from the anterior pituitary. GH stimulates IGF-I production from the liver. This, in turn, has a negative feedback effect on pituitary GH secretion and a positive feedback effect on hypothalamic SRIF release. Pituitary GH secretion has a negative autoregulatory feedback effect on hypothalmic GHRH secretion. GH exerts an effect on somatic growth by a direct as well as indirectly via IGF-I.

gene encoding human Ghrelin has been isolated, and found to be expressed in the stomach and hypothalamus [202]. The isolation of Ghrelin implicates a control system in addition to GHRH/SRIF in the regulation of GH secretion. Ghrelin is analogous to GHRH/SRIF in that it is also synthesized in peripheral tissues (stomach) as well as centrally (hypothalamus). It would be anticipated that hypothalamic Ghrelin would control GH secretion, while peripheral sources may have other biological effects requiring further elucidation. Several artificial GH secretagogues (GHS) release GH in vitro [203]. GHSs, administered alone or in combination with GHRH are potent and reproducible GH releasers and are useful tools for the diagnosis of GH deficiency. GHSs act directly on the pituitary somatrophs [204], stimulating GH secretion by activating the cAmp-dependent protein kinase pathway [205]. GHRH and GHS thus act through distinct receptors, via different intracellular signaling pathways on somatotrope subpopulations. GHS increases the number of somatotropes releasing GH, without altering the amount of hormone released from each individual cell

[206]. Somatostatin too decreases the number of somatotropes that secrete GH, and it is postulated that GHS and SRIF may respectively depolarize and hyperpolarize the same somatotroph cell. GHRH, on the other hand, stimulates the amount of GH secreted per cell, as well as the number of cells secreting GH [206]. Evidence that the main action of GHS occurs at the hypothalamic level, and that the direct pituitary activity is ancillary, is supported by the fact that GHSs require the presence of a functional hypothalamus to exert their GH secreting effect. In patients with intact pituitaries, who have hypothalamic dysfunction leading to stalk section, GHSinduced GH release is blocked and the synergistic effect of GHRH and GHS is absent [207]. Current evidence suggests that the dual control of GH secretion postulated for GHRH/SRIF should be expanded to incorporate Ghrelin. If indeed Ghrelin acts at the hypothalamic level, its mechanism(s) of action could include release of endogenous GHRH. However available data seems to indicate that Ghrelin does not act via GHRH release in that the GH response to GHS is greater than that of GHRH alone [208];

Chapter 3

FIGURE 3.8. A new physiologic model of the regulation of GH secretion involving Ghrelin, GHRH and somatostatin (SRIF). From Diguez and Casanueva [218a].

GHSs potentiate GH release in response to a maximal stimulating dose of exogenous GHRH [209]. GHSs are unable to release GHRH in rat hypothalamic explants [210]; GHRH infusions block GHRH mediated GH release without altering the action of GHS [211,212]; after administration of a saturating dose of GHRh, a second dose of GHRH is ineffective, while GHS is fully effective [213]. The above evidence does not support a role of GHRH in the mechanism of action of GHS. GHSs do not seem to inhibit somatostatin release [212,214–216]. However, GHS mediated GH release is relatively insensitive to the inhibitory action of SRIF [217]. The presence of GHS in hypothalamic structures and the evidence that GHS-elicited GH secretion is not mediated by changes in endogenous GHRH or SRIF, but requires a functional hypothalamus, suggests that exogenous GHS may induce the release of another hypothalamic factor with GH-releasing capability [218] (Fig. 3.8). Release of another unknown hypothalamic factor with GH releasing capability is supposed by observations that GHSs effectively release GH when administered intravenously, subcutaneously, orally or intranasally [219]. Hypoglycemia increases GHS stimulated GH release [214] as does decreased circulating FFAs [220]. GHSs are resistant to wellknown inhibitors of GH secretion including increased FFA levels, hyperglycemia, SRIF infusion, atropine or drugs that enhance hypothalmic SRIF secretion [221]. GHS-mediated GH release is not reduced by a previous rise in GH [213], glucocorticoid treatment [222], or thyroid hormone abnormalities [223]. GHS-mediated GH release is more potent and reproducible than that elicited by GHRH alone [224].

Growth Hormone

89

GHRH-mediated GH release is more erratic and less reproducible, possibly due to the influence of metabolic and hormonal factors. Functional GHs receptors are detected in the human pituitary by the fifth week of gestation [225]. GHS mediated GH-release is demonstratable at birth [221], continues through infancy, increases at puberty and then decreases thereafter. Estrogen and testosterone increase GHS mediated GH release in childhood. When administered with GHRH, GHS has a synergistic action on GH release. As GH secretion elicited by combined GHRH and GHS administration is minimally altered by age, sex, or adiposity and is devoid of potential side effects (unlike insulin-induced hypogylcemia) this test has the potential to become a widely used diagnostic tool in the diagnosis of adult GHD. No side effects have been reported with the GHSs. Although highly specific for GH release, slight increases in prolactin and ACTH/cortisol have been reported with some GHSs [226], leading to the development of new more selective GHSs with no ACTH or PRLreleasing effects e.g. ipamorelin [227]. GHSs are being evaluated as potential therapeutic agents for enhancing circulating GH levels in patients with intact hypothalamic–pituitary axs, including children with GHD as well as adults with catabolic states, AIDS and heart failure. Long-term studies are required with a large number of subjects to determine dosage efficacy.

GH REGULATION

GH Transgenic Animals Transgenesis allows for the transfer of inheritable functional genes between animals of different species. GH transgene models of endocrine hyperfunction have also been useful in investigating regulatory factors required for tissue-specific gene targeting. When a rat or hGH reporter gene driven by a potent metallothionein-I (MT) promoter is expressed in a transgenic progeny, rat or human GH is produced by a number of tissues including the pituitary [228,229]. Circulating GH levels in these animals are several hundred times higher than normal with an increase in transgenic mouse size to twice that of normal littermates [230]. Circulating IGF-I levels are elevated two- to threefold, consistent with the notion that IGF-I mediates the allometric growth of these transgenic animals [228,229]. Interestingly, the effects of excess GH are only apparent after 3 weeks of age and no further growth occurs after 3 months. Immunocytochemical and ultrastructural study of transgenic pituitary glands revealed a marked reduction in both size and number of GH-secreting cells [229,231]. Pituitary lactotroph cells were also suppressed in male transgenic animals, but not in females. In contrast, both sexes exhibited corticotroph cell hyperplasia. Male transgenics had high levels of LH and highly active gonadotrophs, resembling “gonadectomy” pituitary cells. When ovariectomized female

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hGH transgenics were studied, adult female mice developed hypoprolactinemia, but had high levels of LH due to PRL-like activity of GH [232]. Metallothionein rat GH (MTrGH), metallothionein bovine GH (MTbGH) or metallothionein human GH (MThGH) female transgenics are sterile; however, metallothionein ovine GH (MToGH) female transgenics are fertile [233,234]. In MTrGH transgenic animals [235] both the liver and spleen undergo a threefold increase in size. Furthermore, MToGH male transgenics contain hepatic intranuclear inclusions whose presence correlates with high circulating levels of oGH [233]. The heart, lungs, kidneys, and thyroid glands of GH transgenics grew significantly larger when compared to nontransgenic control littermates. Interestingly, no increase was observed in brain or adrenal size. In contrast, methallothionein-driven IGF-I gene (MTIGF-1) transgenics [230] have an increased growth ratio of one to three times that of control littermates, which is only apparent from 6 to 8 weeks after birth. Spleen, pancreas, and kidneys increased in size in direct proportion to the increase in body weight both in IGF-I and GH transgenic models. Compared to GH transgenics, IGF-I transgenic mice show an increase in brain size but no enlargement of the liver or heart [230]. GH transgenics develop renal glomerular sclerosis, similar to abnormalities seen in diabetics, whereas this lesion is not present in IGF-I transgenics. GH transgenics also have elevated serum insulin and lipid levels [236]. Interestingly, mesangial renal proliferation followed by progressive glomerulosclerosis was also seen in transgenic GHRH animals [237], suggesting that the hypersomatotrophism associated with excess GHRH secretion is predominantly GH-mediated. While many of the sequelae of chronic GH overexpression such as increased growth and pituitary somatotroph atrophy were predictable, the abnormalities of hepatic and renal function, and the reduced fertility of female transgenics, were unexpected. These novel findings may provide useful models of human disease. Transgenesis is a useful tool to investigate ontogeny of gene expression, as well as a method of exposing animals to growth at a time when they would not usually be accessible to manipulation of factors responsible for both allometric as well as organ-specific growth. GH Binding Protein (GHBP)

Circulating GH is attached to specific binding proteins [238]. Two circulating GHBPs have been identified, one of high affinity and one of low affinity [239]. Although the site of GHBP synthesis is as yet undetermined, the liver has been postulated as the primary source [240]. The 60-kDa high-affinity BP corresponds to the extracellular domain of the hepatic GH receptor [241]. Under basal conditions, half of the circulating 22-kDa GH is bound to the highaffinity BP when GH levels are up to 10–15 ng/ml [242]. The 20-kDa GH binds preferentially to the low-affinity BP [243]. Binding to plasma GHBP prolongs GH plasma half-

life by decreasing GH metabolic clearance rate, as the GHbinding protein complex is too large for renal glomerular filtration [243]. The high-affinity BP also inhibits GH binding to surface receptors by competing for the ligand [244]. Thus GHBP dampens acute oscillations in serum GH levels caused by pulsatile pituitary GH secretion. Highaffinity BP levels are low in the fetus and neonate, rise most rapidly in the first 1 to 2 years after birth [245,246], and stay constant throughout adult life, with similar levels found in males and females [245]. GH itself does not regulate either type of BP, as evidenced by normal BP levels found in patients with either hypopituitarism or acromegaly [245]. GH resistance, as demonstrated in short stature, Laron dwarfism, and African pygmies, is characterized by decreased plasma levels of high-affinity BP [247,248]. Although these conditions provide insight into the role of GHBP in GH action, Laron dwarfs are also GH receptor-deficient, therefore, no conclusions can be drawn about GHBP function per se from these findings. In African pygmies, GHBP levels, although low, vary widely with considerable overlap with normal subjects.

Peripheral GH Action GH Receptor

GH binds to its peripheral receptor and induces intracellular signaling by a phosphorylation cascade involving the JAK/STAT pathway. GH also acts indirectly by inducing synthesis of IGF-I, the potent growth and differentiation factor [249]. The GH receptor (GHR) is a 70 kd protein member of the class I cytokine/hematopoietin receptor superfamily [250]. GHR consists of an extracellular ligandbinding domain, a single membrane-spanning domain, and a cytoplasmic signaling component. The GH ligand complexes with two GHR components leading to receptor dimerization. This sequential ligand-receptor interaction and dimerization is critical for subsequent GH signaling. GHR dimerization is followed by rapid binding and activation of JAK2 tyrosine kinase, leading to phosphorylation of cytoplasmic signaling molecules, especially the signal transducing activators of transcription proteins (STAT) which comprise the critical signaling components for GH action. These cytoplasmic proteins are phosphorylated by JAK2 and directly translocated to the cell nucleus where they elicit GH-specific target gene expression by binding to nuclear DNA [250]. In addition, STAT1 and STAT5 may in fact interact directly with the GH receptor molecule [251]. Other target actions induced by GH include c-fos induction, IRS-1 phosphorylation, and insulin synthesis. As a differentiating and growth factor, IGF-1 is a critical protein induced by GH, and is likely responsible for most of the growth-promoting activities of GH [252]. IGF-I itself may also directly regulate GH [252] and GH receptor function [253]. The liver contains the highest abundance of GH receptors, and several peripheral tissues also express modest amounts of receptor, including muscle and fat [254].

Chapter 3

Mutations of the GH receptor may be associated with partial or complete GH insensitivity and growth failure. These syndromes are associated with normal or high circulating GH levels, decreased circulating GHBP levels and low levels of circulating IGF-I. Multiple homozygous or heterozygous exonic and intronic GHR mutations have been described, most of which occur in the extracellular ligandbinding domain of the receptor. Tissue responses to GH signaling may be determined by the pattern of GH secretion, rather than the absolute amount of circulating hormone. Thus, linear growth patterns, liver enzyme induction and STAT5b activity may be phenotypically distinct for male animals due to their higher rates of GH pulse frequency [255]. STAT5b is sensitive to repeated pulses of injected GH [256], unlike other GHinduced patterns which are desensitized by repeated GH pulsing. Mice harboring a disrupted STAT5b transgene exhibit impaired male pattern body growth [257] with IGFI and testosterone levels normally seen in female mice. Thus, the sexual dimorphic pattern of GH secretion and GH tissue targeting appears to be determined by STAT5b. The requirement for appropriate GH pulsatility to determine body growth also appears to be mediated by STAT5b [258]. In contrast, STAT5 does not appear to be critical for metabolic effects of GH on carbohydrate metabolism [259]. Unraveling STAT5 regulation in disease, especially those involving GH resistance, will provide novel mechanistic insights for dysregulated GH signaling in humans. Intracellular GH signaling is also abrogated by SOCS proteins, which disrupt the JAK/STAT pathway and thus exert a further level of control over the action of GH [260] (Fig. 3.9).

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91

IGF-I and IGF-II Structure and Synthesis

IGF-I and IGF-II are single-chain polypeptide molecules with three intrachain sulfide bridges (Figs 3.10 and 3.11). IGF-I, composed of 70 amino acids, and IGF-II, consisting of 67 amino acids, have a sequence homology of 62% [261]. The IGFs consist of B and A peptide domains (structurally homologous with the insulin B and A chains), a C domain analogous to the connecting (C) peptide of proinsulin, and a D domain. IGF-I and IGF-II are single distinct gene loci, localized on chromosome 12 (12q22–q24.1) and chromosome 11 (11p15), respectively [262,263]. The IGF-I gene primary transcript can be alternately spliced to different products resulting in IGF-Ia (exons 1,2,3,5) or IGF-Ib (exons 1,2,3,4). Several IGF-I mRNA species have been isolated from adult and fetal tissues. The liver is an important source of circulating IGF-I levels [264]. The IGF-I gene is expressed in human fetal conective tissues and cells of mesenchymal origin [265]. This ubiquitous localization of IGFs favors a paracrine/autocrine function as well as an endocrine function of IGF-I. GH is the major regulator of IGF-I gene expression in adult liver, heart, lung, and pancreas [266], and acts at the level of IGF-I transcription [267]. Fetal IGF-I production is GHindependent [268], and platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF) also stimulate IGF-I production from human fibroblasts in vitro [269]. ACTH, TSH, LH, and FSH stimulate the paracrine production of IGF-I in their respective target tissues. Nutritional status is also an important regulator of IGF-I production at all ages [270]. As discussed below, IGF-I and IGF-II are bound to carrier proteins in the serum. IGFs are found in lymph [271], breast milk [272], saliva [273], and amniotic fluid [274]. IGF-I levels are low before birth, rise during childhood to high levels during puberty, and decline with age [275]. IGF-binding Proteins (IGFBPs)

FIGURE 3.9. Role of STAT proteins in growth hormone signaling. From Herrington, Smith, Schwartz, Carter-Su [260a].

IGF-I and IGF-II are complexed to six specific binding proteins in biological fluids [276] (Fig. 3.12 and Table 3.3). The IGFBPs are cysteine-rich proteins, with very similar amino acid sequences. They have a unique ability to bind IGF’s with high affinity. Recently several groups of cysteine-rich proteins have been described [277] with structural and functional similarities to the IGFBP’s, called IGFBP-related proteins (IGFBP-rPs). The IGFBP-rPs bind IGFs with low affinity. The physiological role of these IGFBP-rPs in the IGF system is not known. However some of these proteins unequivocally bind IGFs [277]. An IGFBP superfamily has been proposed, composed of the IGFBPs and the IGFBPrelated proteins (IGFBP-rPs) (Fig. 3.13). The major form of binding protein present in the human circulation is IGFBPIII, a glycoprotein associated with an IGF molecule and an 80-kDa acid-labile subunit (ALS) to form a 150- to 200kDa complex [278] (Fig. 3.14).

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FIGURE 3.10. Amino acid sequence of human insulin-like growth factor I (IGF-I). The black amino acids are identical to those in human insulin. The numbering corresponds to the numbering of residues in the proinsulin molecule. IGF-I consists of a 70-amino acid single peptide chain with A, B, C, and D, domains. A and B domains are structurally homologous to the A and B chains in the insulin molecule, and the C domain is equivalent to the connecting (C peptide in proinsulin). From Humbel [261a].

FIGURE 3.11. Primary structure of human insulin-like growth factor II (IGF-II). From Sara and Hall [261b].

Actions of the IGFBPs include modulation of IGF actions, storage of IGFs in extracellular matrices, and the carrier protein function of IGFBPs. In the serum most of the IGF circulates as a 150,000 dalton complex that consists of IGF-I or IGF-II plus IGFBP-III and a nonIGF binding component, the acid labile cubunit (ALS). Binding of IGF-I or II to IGFBP-III in the presence of ALS results in the formation of an IGF-IGFBP-ALS ternary complex, which is stabilized by IGF binding [279] 75% of circulating IGF-I and IGF-II is carried in this ternary 150-kDa complex. When associated with the 150-kDa complex, the IGFs do not readily leave the vascular compartment, and have prolonged half-lives of 12 to 15 hours [280] compared to the half-life of free IGF-I which is less than 10 minutes [281]. The exact function of the large store of IGF bound

to the 150-kDa ternary serum complex is unknown. It has been hypothesized to be a functional pool of material available for organisms to use during stress. The plasma concentrations of IGFBPs are hormonally regulated. Serum IGFBP-III levels correlate with IGF-I and II levels, double in patients with acromegaly and are reduced in hypopituitarism [282]. Malnutrition, insulin-dependent diabetes mellitus, and cirrhosis are associated with decreased IGF-I levels, as well as suppressed IGFBP-III levels [283]. IGFBP-I levels are high at birth and decline until puberty [284]. There is a diurnal variation with a nocturnal peak in serum IGFBP-I levels [285]. However, IGFBP-I levels are elevated in hypopituitarism [286] and decreased in acromegaly. IGFBP-I levels are regulated by insulin. The increased IGFBP-I levels associated with insulin-dependent

FIGURE 3.12. Simplified diagram of GH-IGF-I axis involving hypophysiotropic hormones controlling pituitary GH release, circulating GH-binding protein and its GH receptor source, IGF-I and its largely GHdependent binding proteins, and cellular responsiveness to GH and IGF-I interacting with their specific receptors. IGFR, IGF-I receptor; FFA, free fatty acids. From Rosenbloom [276a].

Table 3.3.

General characteristics of the human IGF binding protein No. of amino acids

Core molecular mass (kDa)

Chromosomal localization

IGF affinity

Modulation of IGF action

Source in biological fluids

IGFBP-I

234

25.3

7

I = II

Inhibition and/or potentiation

Amniotic fluid, serum, placenta, endometrium, milk, urine, synovial fluid, interstitial fluid and seminal fluid

IGFBP-II

289

31.4

2

II > I

Inhibition

CSF, serum, milk, urine, synovial fluid, interstitial fluid, lymph follicular fluid, seminal fluid, and amniotic fluid

IGFBP-III

264

28.7

7

I = II

Inhibition and/or potentiation

Serum, follicular fluid, milk, urine, CSF, amniotic fluid, synovial fluid, interstitial fluid, and seminal fluid

IGFBP-IV

237

25.9

17

I = II

Inhibition

Serum follicular fluid, seminal fluid, interstitial fluid and synovial fluid

IGFBP-V

252

28.5

5

II > I

Potentiation

Serum and CSF

IGFBP-VI

216

22.8

12

II > I

Inhibition

CSF, serum and amniotic fluid

Modified from Rajaram S et al. [278a].

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FIGURE 3.13. The IGF system. The components of this system include the peptides IGF-I and -II, IGFBP-1 to -6, IGFBP-rPs, IGFBP proteases, type I and type II IGF receptors, and potential IGFBP(s) and IGFBP-rP(s) receptors. M6P, Mannose-6-phosphate. From Hwa, Oh, Rosenfeld [277a].

diabetes mellitus [287] are normalized by insulin, and insulinoma is associated with suppressed IGFBP-I levels. Octreotide (a long-acting somatostatin analog) has been shown to increase IGFBP-I levels in acromegalic patients [288]. When normal subjects ingest glucose, the fall in IGFBP-I levels correlates inversely with the rise in insulin levels [289]. Hypophysectomy is associated with elevated IGFBP-II levels in rats, which fall with GH administration [283]. Insulin infused into normal rats increases IGFBP-II levels [283]. PARACRINE GH ACTION GH and GH-releasing factors are also produced in tissues outside the hypothalamic pituitary axis. The extrapituitary actions of GH are likely autocrine/paracrine and complement the classic endocrine action between the GHreleasing factors, GH and target tissues. GH gene expression is not restricted to the pituitary gland. GH immunoreactivity and GH mRNA have been localized to several extrapi-

tuitary tissues [290] including placenta, mammary gland muscle, spleen lymphocyte, suggesting an extrapituitary paracrine/autocrine action for GH. Classic endocrine actions of GH include GH promoting effects on bone and several metabolic actions.

Effect on Growth Whether GH exerts its growth-promoting action on the skeleton via a direct GH effect on the growth plate, or indirectly by stimulating hepatic or local IGF-I production, remains unclear. The somatomedin hypothesis proposed an indirect effect of GH on growth mediated through plasmatransported IGF-I that is synthesized in the liver [291]. The dual effector theory postulates a direct effect of GH by stimulating epiphyseal growth plate precursor cells to differentiate and become responsive to IGF-I, and to induce local IGF-I synthesis that in turn stimulates the multiplication of differentiating chondrocytes [292]. This proposes a dual role for GH and IGF-I in the control of tissue mass. IGF-1 has

Chapter 3

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95

which is rapidly restored by GH administration [299]. Hypophysectomized rats show a progressive decline in longitudinal bone growth rate [300], finally growing at only 0 to 10% of the normal rate. GH and IGF-I Effects on Bone in vitro

FIGURE 3.14. Relative distribution of various IGF pools in human serum. The distribution of IGFs between the 50-kDa, 150kDa, and the free pool as determined before and during continuous subcutaneous infusion of 30 mg/day of rhIGF-1 in healthy men. From Rajaram, Baylink, Mohan [278a].

a potent mitogenic action, especially on immature differentiating cells (adipocytes, chondrocytes), whereas GH is not a mitogen [293]. The dual effector theory proposes that although the overall effects of GH and IGF-I on growth at the tissue level appear to be the same, at the cellular level their effects are very different; GH has both a direct action and an indirect action mediated by IGF-I. The IGFBP regulatory hypothesis proposes that GH stimulates both IGF-I and IGFBP-III [294]. Circulating IGFBP-III regulates the bioavailability of IGF-I to the growth plate, and may inhibit IGF-I action. However, recently hepatic IGF-II production in the liver of mice was completely abolished using an IGF-I knockout mouse model, with complete inactivation of the IGF-I gene in the hepatocytes [295]. Serum IGF-I concentration was reduced by 75%. However the reduction in serum IGFI concentration had no discernible effect on postnatal body growth. Role of GH in Growth and Development

Congenital absence of the human pituitary gland does not result in abnormal birth or newborn weight [296], indicating that fetal and early postnatal development may occur independently of GH. Postpubertal mammalian longitudinal bone growth results from chondrocyte proliferation and differentiation in long bone epiphyseal growth plates. The proximal zone of the epiphyseal growth plate contains stem cells and is the layer of prechondrocyte origin [292]. These cells differentiate and enter the proliferative layer, where they undergo limited clonal expansion. After maturation, the cells become calcified and are incorporated into metaphyseal bone [297]. Hypophysectomy in the growing rat results in a marked reduction in tibial epiphyseal cartilage plate width [298],

Although earlier studies failed to demonstrate in vitro stimulatory effects of GH [301], subsequent studies have demonstrated significant direct effects of GH on chondrocyte and osteoblast cultures [302]. GH increases progenitor cell thymidine incorporation, chondrocyte differentiation, and osteoclast formation. The in vitro effect of IGF-I on chondrocytes has been more consistently demonstrated. IGF-I increases thymidine and sulfate incorporation into isolated chondrocytes [303], and this effect is additive with GH [302,303] occurring at different stages of maturation [304]. Cells from the proximal growth plate (prechondrocytes and young proliferating chondrocytes) form larger colonies in vitro, whereas chondrocytes from the more distal proliferative zone form smaller colonies [305]. Both GH and IGF-I induce chondrocyte colony formation [305] but exert differential effects on colonies of different sizes. GH preferentially potentiates formation of large size colonies, suggesting that GH stimulates prechondrocyte differentiation [304], while IGF-I induces smaller colonies, implicating IGF-I in epiphyseal chrondrocyte stimulation at a more mature stage of differentiation [304]. In vivo Effects of GH and IGF on Skeletal Growth

GH administered locally at the epiphyseal growth plate [306,307] or infused into a femoral artery [308] in hypophysectomized rats results in unilateral longitudinal bone growth and an increase in growth plate width. When IGFI antiserum is infused locally with GH this growth effect is eliminated [308], suggesting that GH exerts its growth promotion via local IGF-I. Local infusion of IGF-I alone into the epiphyseal growth plate results in longitudinal bone growth [307] and increased epiphyseal width. Local GH infusion at the growth plate additionally results in increased numbers of IGF-I-containing cells in the proliferative zone, as well as producing a simultaneous increase in plasma IGFI levels [309], suggesting that the in vivo GH effect may be mediated by locally produced IGF-I in cells that have been stimulated by GH. This may explain why GH-deficient children treated with GH grow significantly, even though their plasma IGF-I levels may not change appreciably [310]. The effects of systemic GH and IGF-I administration have been compared. IGF-I increases tibial epiphyseal width [311] and total body growth in hypophysectomized rats [312], but to a lesser degree than after replacement doses of GH. In mutant GH-deficient dwarf rats, hGH infusions had a more pronounced stimulatory effect on body weight and bone growth than hIGF-I administration, while hIGF-I caused a greater increase in kidney, adrenal, and spleen size than did hGH [313]. GH and IGF-I may also have

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differential tissue effects. Some of these observations suggest that GH acts directly on tibial plate cartilage via a locally produced paracrine or autocrine IGF-I action. The somatomedin hypothesis, however, suggests that plasma IGF-I levels are much higher than tissue levels, and that plasma IGF-I is indeed biologically active. The liver synthesizes more IGF-I in response to GH stimulation than does costal cartilage. Furthermore, IGF-I increases thymidine and sulfate uptake and incorporation in bone and cartilage cultures to a far greater degree than does GH. These findings favor the liver as the predominant site of IGF-I synthesis, with subsequent plasma transport of biologically active IGF-I to its target organ, bone, where it mediates growth. A model incorporating GH, IGF and IGFBP systems in the regulation of systemic growth integrates the dual effector theory and the somatomedin hypothesis (Fig. 3.15). GH stimulates hepatic/tissue IGF-I synthesis, and it is the endocrine action of this circulating IGF-I on the growth plate which regulates skeletal growth, by stimulating cell division. Furthermore, GH induces hepatic IGFBIII and ALS production, forming the circulating 150-kDa ternary

complex that binds IGF-I, thereby increasing its half-life. GH also directly induces prechondrocyte differentiation into chondrocytes at the growth plate, in addition to stimulating local IGF-I production, which acts in an autocrine/ paracrine fashion to further stimulate cell division. Thus the endocrine actions of circulating IGF-I largely contribute to cell division which results in intrauterine and post natal growth. This mechanism is augmented by the direct actions of GH stimulating prechondrocyte/chondrocyte differentiation at the growth plate as well as the local IGF-I stimulation by GH where it acts by an autocrine/ paracrine action to stimulate cell division.

GH and Metabolism In childhood, the multiple metabolic actions of GH contribute to linear growth. GH continues to be secreted in adulthood after growth cessation, implying that GH has important metabolic functions in adult life. The metabolic actions of GH are either acute insulin-like or chronic insulin antagonism. The metabolic actions of GH may be direct or indirectly mediated by IGF-I.

FIGURE 3.15. Integrated model of the GH-IGFBP-IGF axis in the growth process. Three mechanisms are proposed: (1) GH stimulates IGF-I production; circulating IGF-I (endocrine IGF-I) acts at the growth plate; (2) GH regulates hepatic production of IGFBP-III and ALS: IGF-I binds to IGFBP-III and thereafter with ALS, forming the 150 kDa ternary complex; proteases cleave IGFBP-III into fragments that release IGF-I in the intravascular space and at the growth plate; and (3) GH induces differentiation local IGF-I production, and IGF-I acts via an autocrine and paracrine mechanism to stimulate cell division.From Spagnol and Rosenfeld [313a].

Chapter 3 Table 3.4.

Acute metabolic effects of growth hormone

Enhanced lipolysis and lipid oxidation ÆMobilization of stored triglyceride Protein synthesis stimulation Antagonism of insulin action Water and sodium retention

Acute, transient, insulin-like effects of GH can be demonstrated on carbohydrate, protein, and lipid metabolism (Table 3.4). The insulin-like effects are more easily demonstrated in tissues that have not recently been exposed to GH (for example, those derived from hypophysectomized animals, or GH-deficient humans) than in normal tissues [314]; casting doubt on the physiologic relevance of the insulin-like effects of GH, because GH is secreted endogenously in frequent bursts, resulting in significant changes in ambient GH levels. The chronic antiinsulin effects of GH result in decreased glucose utilization, increased lipolysis, and tissue refractoriness to the insulin-like effects of GH. Antiinsulin effects occur at both hepatic and peripheral sites [315]. Effects on Carbohydrate Metabolism

GH-deficient children have decreased fasting glucose levels [316,317], decreased insulin secretion [316], contradictory impairment of glucose tolerance [318–321], and increased insulin sensitivity due to increased glucose utilization and blunted hepatic glucose release. GH replacement increases fasting glucose levels [317,318], insulin levels [318], and hepatic glucose production [317]. Endogenous physiologic GH secretion antagonizes insulin action [322]. Normally, GH secretion increases 3 to 5 hours after oral glucose ingestion, resulting in decreased disposal of a second oral glucose challenge, associated with hyperinsulinemia occurring 2 hours after GH levels peak. Both intravenous and oral glucose tolerance tests are impaired if performed during periods of increased GH secretion, such as sleep onset. GH-deficient adults have elevated fasting insulin levels and a positive correlation between fasting plasma insulin and both fat mass and waist girth [323], suggesting insulin resistance, which is confirmed by hyperinsulinemic euglycemic clamp [324]. GH replacement initially further increases insulin resistance, in the first 1 to 6 weeks of therapy, but subsequently, although hyperinsulinemia persists, carbohydrate metabolism returns to baseline after 3 months of GH treatment [325]. Acromegaly is the experiment of nature for effects of excess GH secretion on carbohydrate metabolism. Several stages of impaired glucose tolerance occur in acromegaly, possibly indicative of a stepwise deterioration in carbohydrate metabolism as the disease progresses [326]. Glucose tolerance after both intravenous [326] and oral glucose administration is impaired in association with hyperinsu-

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linemia [327]. When acromegalic patients eventually manifest overt diabetes mellitus, postglucose insulin levels are depressed [327]. Effects on Protein Metabolism

Somatic growth is under the primary control of GH and, as such, GH is an anabolic hormone. GH causes urinary nitrogen retention and decreased plasma urea levels in GH deficient children [328], and in normal [329] and obese [330] adults. Furthermore, GH-deficient children increase their muscle mass following GH treatment [331]. Both insulin and IGF-I have been implicated in the anabolic effects of GH on protein metabolism, especially as insulin is a protein anabolic hormone and GH stimulates circulating insulin levels. However, GH-stimulated protein synthesis precedes the GH stimulated increase in insulin levels. Physiologically, GH may promote nitrogen retention during starvation, as GH levels rise during fasting.

Effects on Lipid Metabolism GH Effect on Adipose Tissue

GH plays an important role in fat metabolism. GHdeficient children are mildly obese [331,332], with a decreased total number of fat cells [333,334] that are larger in size, with a greater lipid content than normal. GH replacement therapy leads to a decrease in body fat and, eventually, a decrease in the size and lipid content of subcutaneous adipocytes. GH deficient adults have altered body composition, with increased fat mass and decreased lean body mass. Initial acute effects of GH on lipid metabolism are antilipolytic (insulin-like) and subsequently, GH exerts a lipolytic (antiinsulin) effect. Lipolysis

GH increases fat mobilization and decreases fat deposition, thereby decreasing body fat. GH activates hormonesensitive lipase [335], resulting in increased hydrolysis of triglycerides to free fatty acids and glycerol (lipolysis). GH also decreases fatty acid reesterification [336]. The lipolytic actions of GH can be counteracted by insulin and are additive with glucocorticoids [337], which decrease the antilipolytic effects of insulin [338]. In vivo Effects on Lipids and Lipoproteins

Physiologic doses of GH have multiple, inconsistently demonstrated effects on free fatty acids and ketones [339]. Lipolytic and ketogenic effects of GH are easily demonstrable in normal subjects with suppressed insulin levels [318] and in GH-deficient insulin-dependent diabetic patients infused with GH [340,341]. GH deficient adults have elevated total cholesterol, low density lipoprotein cholesterol (LDL) and apolipoprotein B (ApoB) [342], with low high density lipoprotein levels (HDL) and high triglyceride levels. This lipid profile is

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associated with premature atherosclerosis and cardiovascular disease. GH replacement treatment, decreases total cholesterol (342) LDL cholesterol and ApoB and increases HDL. Long term surveillance will be required to determine whether GH replacement therapy reverses premature atheroclerosis and reduces cardiovascular morbidity and mortality in GH-deficient adults.

the abdomen [323]. The reduction in abdominal fat mass is mainly due to reduced visceral fat mass [343,344]. GH therapy increases total body water [343] especially extracellular water within 3 to 5 days [345,351]. Total blood volume increases after 3 months of treatment [346]. A direct action of GH and/or IGF-I on renal tubular sodium reabsorption, as well as activation of the renin angiotensin system [345,352], contributes to the antinatiuretic GH effects.

GH Effects on Body Composition

The anabolic, lipolytic and antinatiuretic actions of GH impact body composition affecting fat mass, lean body mass and fluid volume in GH-deficient adults. Lean body mass (LBM) is reduced [323,343], and fat mass is increased in GH-deficient adults compared to predicted values for age, sex, and height matched normal controls in GH-deficient adults [323]. Excess fat accumulates mostly in the visceral component in a central distribution, mainly abdominal [343,344]. Total body water is reduced in adults with GH deficiency. Reduced plasma volume [345] and total blood volume [346], contribute to the reduced extracellular water [347,345]. GH replacement therapy reverses these effects on body composition by increasing LBM (Fig. 3.16) [343,344, 348–350]. There is a significant increase in skeletal muscle, the most important component of LBM [348]. GH replacement also reduces fat mass by 4–6 kg in GH deficient adults [343,344,348–350] with the most significant reduction in

GH Effects on Bone Metabolism and Bone Mineral Density (BMD)

GH-deficient adults have reduced bone mass, [353] measured by bone mineral density (BMD), resulting in a 2 to 3 times increased fracture rate [354]. The pathogenesis of the bone loss is unclear, with studies demonstrating unchanged [355], increased, [356] or reduced [357] bone formation as well as unchanged bone resorption [358] and delayed mineralization [359]. With GH replacement therapy there is predominant bone resorption in the first 3 to 6 months, resulting in initial reduced BMD after 6 to 12 months [360] with subsequent increased bone formation leading to a net gain in bone mass after 18 to 24 months of treatment [361]. GH Effects on Muscle Strength and Exercise Performance

GH deficiency is associated with reduced muscle strength, due to altered body composition [362]. Reduction in muscle cross-sectional area, as well as lack of conditioning and training, may contribute to the weakness. Prolonged GH replacement therapy for at least 12 months is required to significantly increase muscle strength [362] and for at least 3 years to normalize the muscle strength [363]. GH Effects on Cardiovascular Function

FIGURE 3.16. Effect of GH replacement on (a) lean body mass (LBM) and (b) fat mass in 24 adults with GHD. Reproduced with permission from Salomon et al. [347a].

Hypopituitary adults have increased vascular mortality [364] mostly attributable to cerebrovascular and cardiovascular disease [365]. GH deficiency has been implicated [366] based on the hypothesis that adult GH deficiency predisposes to the development of premature atherosclerosis, as evidenced by increased carotid artery intima-medial thickening and plaque formation [367], increased triglycerides, plasminogen activator inhibitor-1 activity and fibrinogen [368]. The role of GH in regulating cardiac structure and function has not been established. In GH-sufficient adults with dilated cardiomyopathy and congestive heart failure, GH has been shown to improve cardiac function and increase ventricular mass [369] by some workers and to have no beneficial effect by others. GH deficient adults have reduced left ventricular mass and impaired systolic function. Short term GH treatment for 6 months improves the deficits in cardiac function and structure, with a sustained effect on cardiac performance for up to 3 years of GH treatment [370]. Effects of GH excess in acromegaly manifest as left ventricular hypertrophy, hypertension, cardiac failure and increased mortality due to cardiovascular causes [371]. The long term effects of GH

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replacement on cardiac structure and function are unknown. GH replacement should aim to restore somatic deficiency, including cardiac size and function. In view of the deleterious cardiac effects of excessive GH, supraphysiological GH doses should be avoided.

surements to be consistently more reproducible and sensitive than repeated pharmacologic stimulation tests [377]. These conflicting findings may be attributable to differences in control groups in the discrepant studies.

TESTS OF GH SECRETION

An alternative approach to be assessment of spontaneous GH secretion is the measurement of GH concentrations in urine [378]. As urinary GH concentrations are very low, assays require high affinity antibodies [379]. Several different immunoassay methods for urinary GH measurement have been developed [380], none of which have the ability to reflect pharmacological GH testing or to adequately discriminate between normal and abnormal GH secretion [381–383]. Clinical utility of urinary GH measurements in the diagnosis of GH deficiency will require attainment of adequate-age-and-sex-matched controls and a decision as to whether GH excretion should be expressed relative to body weight or creatinine excretion [384]. 24-hour urinary collections have been assayed for GH in an attempt to diagnose acromegaly [385,386]. Overnight urinary GH measurements were, however, not useful in differentiating acromegalic patients from normal controls [385]. However, others have found 24-hour urine collections for GH measurement to be useful in diagnosis of acromegaly [385].

Urinary GH Measurement

Because of the pulsatile nature of pituitary GH secretion, a single random blood sample for GH measurement is not helpful in the diagnosis of GH hypersecretory or deficiency states, or GH neurosecretory disorders. Nonphysiologic provocative or suppression tests, or measurement of spontaneous GH secretion by 24-hour integrated serum GH concentration (IC-GH), are therefore employed to assess GH secretion.

Measurement of Spontaneous GH Secretion Twenty-four-hour Integrated Concentration of GH

Pituitary GH secretion occurs episodically during waking hours, as well as during sleep, necessitating continuous measurement of integrated secretion over 24 hours [372] rather than over shorter periods. Constant blood collection over a 24-hour period allows the determination of a true mean or IC-GH. Measurement of the 24-hour IC-GH concentration ideally requires a nonthrombogenic continuous withdrawal pump [373] or patent indwelling catheter from subjects whose food intake and physical activity are not limited. Sampling intervals of 20 minutes are most widely used [373], but 5-minute and 30-second sampling frequencies detect significantly more pulses per hour [374]. Samples from collection periods may be pooled, producing a combined aliquot in which the ICGH concentration is measured. The 24-hour IC-GH reflects the average GH concentration over a 24-hour period, eliminating peak or trough levels that might otherwise be obtained by single random sampling of GH when the latter is released in a pulsatile manner. The discriminating power of continuous GH measurement in the diagnosis of GH deficiency has been disputed [375,376]. Measuring spontaneous GH secretion in prepubertal short children was found to be an insensitive test, with no clear diagnostic advantage over GH stimulation tests and considerable overlap between values obtained in normal, short children and children with GHD [375]. As young normal controls have IC-GH levels which overlap with those of organic hypopituitary patients [377] measurement of spontaneous GH secretion has limitations in the diagnosis of organic GH deficiency in adults. As measurement of spontaneous GH secretion has limitations in the diagnosis of adult GH deficiency, 12-hour overnight spontaneous GH secretion has been measured. Night-to-night variation in results occurred when the test was repeated on consecutive nights, as compared with GH stimulation tests which proved to be more reproducible [376]. However, others have found 24-hour IC–GH mea-

Evaluation of GH Hypersecretion Increased serum IGF-I levels are a consistent finding in acromegaly [387]. Integrated 24-hour serum GH levels are elevated and show a log (dose) response correlation with serum IGF-I levels [388]. The currently accepted diagnostic test of GH hypersecretion is failure of GH levels to be suppressed to less than 1 ng/mL within 2 hours following a 75 g oral glucose load using an IRMA (two site immunoradiometric assay) or chemiluminescent assay [389]. In normal subjects receiving oral glucose loading, serum GH levels initially fall and then subsequently increase as plasma glucose declines. However, in acromegalic patients, oral glucose fails to suppress GH to the normal range [390]. GH levels may increase in response to an oral glucose load in acromegaly (in approximately 28%) [391], remain unchanged (in approximately 36%), or fall (in approximately 36% of patients). As GH secretion is pulsatile, random GH measurements may be misleading.

Evaluation of GH Deficiency Random GH and 1GF-I Measurements

Random GH measurements are not diagnostic of GH deficiency, as GH secretion is pulsatile and daytime levels are often low in normal subjects. Low IGF-I levels are suggestive of GH deficiency, but are also found in malnutrition [392], acute illness, celiac disease, poorly controlled diabetes mellitus, liver disease, and estrogen ingestion [393]. Fifteen

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percent of children diagnosed to be GH deficient by stimulation tests may have normal IGF-I levels [394]. IGF-I levels are normally very low before 3 years of age and highest in adolescence. Normal and GH-deficient children may have IGF-I levels which overlap with those observed in infancy [395]. Furthermore, both normal and low IGFI levels are present in children with growth delay and genetic short stature [396]. IGF-I levels do not always correlate with GH levels after provocative GH stimulation [397]. Low IGFBP-III levels are also encountered in patients with GH deficiency [398]. The diagnosis of adult GHD is more reliably established by provocative testing of GH secretion, as a single ageadjusted IGF-I level has limited diagnostic value [377]. Whereas, in acromegaly there is an excellent correlation between IC-GH and IGF-I [388], in adult GH deficiency, age-adjusted IGF-I should be regarded as a simple, low-cost screening test, because of the limitations outlined above. IGFBP-III is the major serum carrier protein for the IGF peptides, and is GH-dependent [282]. IGFBP-III assays have several advantages over IGF-I assays in that (i) they are technically simple, not requiring separation of IGF peptides from BPs; (ii) circulate in the plasma in high concentrations, so that assay sensitivity is not compromised; (iii) IGFBP-III plasma concentrations are less influenced by nutritional factors than IGF-I levels; (iv) plasma levels vary minimally with age, so that identification of abnormal concentrations in infancy or puberty is clear-cut [384,399]. IGFBP-III is a sensitive test for diagnosis of severe childhood GHD [518], with superior reproducibility to GH stimulation tests [400]. Provocative Tests

Dynamic testing of GH reserve involves stimulation of the somatotrophs to secrete GH in response to a pharmacologic stimulus. Several GH stimulatory agents have been utilized, including insulin, clonidine, arginine, L-dopa, GHRH, and propranolol. Insulin-induced Hypoglycemia (Insulin Tolerance Test, ITT)

This reliable stimulus for GH secretion is the gold standard provocative test [401]. The ITT is the best validated GH stimulation test and is recommended by the Growth Hormone Research Society as the test of choice [401]. Regular insulin 0.1 IU/kg is administered intravenously to decrease the basal glucose level by 50% to a value below 40 mg/dI. Maximal GH secretion peaks at 30 to 60 minutes. Patients may experience symptoms of hypoglycemia, including light headedness, anxiety, tremulousness, sweating, tachycardia seizures and rarely, unconsciousness. Insulin induced hypoglycemia is contraindicated in patients with a history of seizure disorder, coronary artery disease, or over the age of 55 years. The test should be performed under close supervision, and intravenous glucose (50%) should be readily at hand for rapid administration. The risk of inducing profound hypoglycemia is greater in GH-deficient patients because of their increased sensitivity to insulin.

Clonidine

This alpha-adrenergic agonist stimulates GH release via a central action [402]. Clonidine (0.15 mg/m2) is administered orally, with a maximum GH secretory peak occurring after 60 to 90 minutes. Patients may experience some drowsiness, with a decrease in systolic blood pressure in sodium depleted GH deficient adults at doses required to release GH (0.25–0.30 mg orally.) Clonidine, frequently used as a stimulus for GH release in children, is not reliable in adults [403]. L-dopa/propranolol

l-dopa, the immediate metabolic precursor of dopamine, stimulates GH release by stimulating hypothalamic dopaminergic receptors. Adrenergic blockade (propranolol) enhances the GH response to l-dopa [404]. l-dopa is administered orally according to the patient’s weight (125 mg if weighing <10 kg; 250 mg if 10–30 kg; 500 mg if >30 kg) together with propranolol 0.75 mg/kg (maximum dose 40 mg) after an overnight fast. Maximum GH secretion occurs after 60 to 90 minutes. l-dopa is very effective in stimulating GH release and rarely results in adverse effects [255]. Arginine/GHRH

Recently it has been shown that arginine potentiates maximal somatotroph responsiveness to GHRH [405]. After an overnight fast, GHRH 1 ug/kg is administered as an intravenous bolus at 0 minutes with Arginine 30 g in 100 ml infused from 0 to 30 minutes, with subsequent blood sampling for GH performed every 15 minutes for 90 minutes. Combined Arginine/GHRH responses are ageindependent and is a highly reproducible GH provocative test [405], at least as sensitive as insulin induced hypoglycemia [406]. GHRH/GHRP-6

GHRH 1 ug/kg plus GHRP-6 1 ug/kg is given intravenously at 0 minutes and blood is drawn for GH sampling at 0 and 120 minutes [407]. GH releasing peptide-6 (GHRP-6) is an artificial hexapeptide [203] that activates a hypothalmic and pituitary receptor whose natural ligand Ghrelin was recently characterized [202]. Combined administration of GHRP-6 and GHRH is the most potent stimulus to GH release, with excellent reproducibility and no side effects [203]. GHRH/GHRP-6 is a good alternative to the ITT in patients with organic pituitary disease, however there is some overlap between GH levels attained in the control group and severely GH-deficient patients. Since GHRH and GHRP act directly on the pituitary, it is possible that their administration restored GH secretion in patients who had a deficiency of these secretagogues because of hypothalamic disease [408]. Pitfalls of Provocative Testing

Provocative testing has several limitations. The diagnosis of childhood onset GH deficiency is determined by an inad-

Chapter 3

equate GH response to two distinct provocative tests. The definition of an adequate response is arbitrary. The availability of recombinant human GH in the mid-1980s resulted in a loosening of the diagnostic cut-off from 5–7 ng/ml to 10 ng/ml. Sequential tests on the same day have been attempted [409], but discordant individual results are obtained in approximately 30% of subjects. By altering the temporal relationship between the administration of provocative agents, the amount of GH released can be decreased or increased [410–412]. A combination of two provocative agents administered simultaneously has been tried, but this approach failed to identify some patients who demonstrated GH deficiency following separate testing with each agent individually [413]. The reproducibility of provocative tests has been questioned, as only a moderate correlation was found between pharmacologic tests carried out on two separate occasions, 6 weeks apart [414]. Many physical states alter stimulated GH levels. Increased GH response to various provocative stimuli during puberty [415] have been attributed to increased estrogens and androgens [416]. Sex steroid priming with either estrogens or testosterone 3 to 5 days prior to provocative testing in prepubertal children has therefore been recommended by some. Mildly impaired GH release in response to provocative stimuli in hypothyroid patients is a frequent finding [417]. Patients should thus be euthyroid at the time of provacative GH testing. Furthermore, 67% of patients with “idiopathic” GHD diagnosed in childhood normalize their GH secretion, when reevaluated after completion of treatment [418].

Variability of GH Assays The comparisons of results of various GH assays obtained in different laboratories is difficult because of differences between almost all aspects of the immunoassays. Plasma GH is usually measured by radioimmunoassay (polyclonal or monoclonal; RIA) or by immunoradiometric assay (dual monoclonal; IRMA). Comparative GH measurements are obtained using 11 commercial immunoassays varied by a factor of three [419]. As previously mentioned, there are several different circulating forms of GH, most of which are not measured in GH assays because reagents to detect all these forms do not exist. Because monomeric 22 k is the only GH form available as a standard in sufficient purity and quantity, and because monomeric 22 k is also the most abundant circulating form, it is used as the basis for GH measurement: however it accounts for only about 25% of circulating immunoreactivity [420]. The other GH forms are recognized to varying and largely unknown degrees. Thus an assay result is an averaged look at a variety of GH forms, and different antibodies or assay designs yield different answers. Polyclonal antibodies were used in the early RIAs, inducing higher estimates of GH, because they recognized several molecular forms of GH compared to newer immunometric

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assays employing highly specific monoclonal antibodies [421,422]. GH standards also affect comparison of GH values in different laboratories. In 1994, the first international standard for somatotropin, IRP 88/624 [423], was prepared by the WHO, using recombinant technology in contrast to the previous standards prepared from pituitary extracts. Use of an international standard enables uniformity of calibration between different GH kits. GH-binding proteins (GHBPs) may influence GH estimates by interfering in some of the GH assays. In the circulation, approximately 50% of GH is complexed to GHBP, which corresponds to the extracellular domain of the GH receptor. Both high affinity and low-affinity BPs are present in the serum. As the low-affinity GHBP has a greater affinity for the 20-KD GH molecule, it presumably does not interfere in GH estimates in the new GH assays specific for the 22-KD GH molecule. The influence of GHBP in noncompetitive immunometric assays could lead to too low estimates of GH if the GHBP competes with one or both antibodies. In competitive assays like the RIAs, the error in the GH estimates will depend on whether the GHBPbound 125 I-GH is in the antibody-bound or in the free fraction. In RIAs using polyethylene glycol or double antibody for separation, the GHBP-complexed GH will end up in the free fraction if the antibodies are directed against epitopes in the GHBP-binding area and thus give spuriously high values [424]. Another factor contributing to the heterogeneity of results from GH immunoassays is the antibody used. As different antibodies employed in immunoassays bind to a different spectrum of GH isoforms, GH concentrations measured by immunoassay are likely to depend on the particular antibody used. Furthermore, as the distribution of the different GH isoforms varies between individuals, results from different GH immunoassays cannot be compared. The heterogeneity of GH immunoassay results poses a major problem in the definition of standards for the diagnosis of GHD. Consensus statements including cut-off recommendations refer to the polyclonal-antibody-based RIA. However RIA is now used infrequently, therefore clinicians should be aware of the GH assay used by their laboratory and how it compares to polyconal RIA. Recently, three new GH assays have been developed, two of which, ESTA and IFA, measure GH bioactivity rather than immunoactivity as measured by convential RIA. A third, GHEA, measures the different GH isoforms in the circulation [425]. ESTA or IFA bioassays could replace RIA if they become commercially available, as they more closely reflect biologically active GH than that measured by RIA. CLINICAL USE OF GH

Recombinant hGH (rhGH) The application of recombinant DNA technology has made available a sufficient quantity of hGH for treatment of GH

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deficiency [426]. Early trials investigating the effects of recombinant human GH compared to pituitary-derived hGH in adults showed comparable kinetics in the two preparations, as well as similar ability to stimulate IGF-I levels. Both compounds resulted in a similar decrease in serum blood-urea nitrogen (BUN) and cholesterol, and increased serum triglyceride levels [427] and clinical growth-promoting effects.

GH Therapy in Childhood Classic GH Deficiency

Childhood GH deficiency ranges from complete absence of GH associated with severe growth retardation to partial GH deficiency resulting in short stature. Diagnosis is based on decreased height (more than 2.5 standard deviations (SD) below the mean height for age-matched normal children), poor growth velocity (less than the twenty-fifth percentile), delayed bone age and a predicted adult height below mean parental height [428]. GH deficiency (GHD) is usually confirmed by inadequate pituitary GH responses to standard provocative stimuli (peak serum GH value of less then 10 mg/L) [429], with values less than 5 mg/L in severe GH deficiency. Combined clinical evaluation and provocative testing are used in the assessment of childhood GH deficiency. Concomitant endocrine deficiencies, especially hypothyroidism, should be corrected to maximize the growth-promoting benefits of hGH. GH replacement should be started as early as possible before height drops below the third percentile, as total height gain is inversely proportional to the pretreatment chronologic and bone age [430], as well as severity of GH deficiency. The most pronounced acceleration in linear growth rate occurs during the first two years of treatment [431]. Dose and frequency of administration of hGH both influence height velocity [432]. Currently, the recommended weekly dose is 0.18 mg/kg (0.54 iu/kg) divided into six or seven daily subcutaneous injections or as three injections/ weekly administered on alternate days. The maximal replacement weekly dosage is 0.3 mg/kg (0.9 iu/kg) divided into equal doses given on three alternate days. Idiopathic Short Stature

Idiopathic short stature describes otherwise normal children who are at or below the fifth percentile for height, with normal growth hormone responses to provocative stimuli. In the past, scarcity of human pituitary GH limited its therapeutic uses to children with severe GH deficiency diagnosed by a decreased growth rate and inadequate GH response to two provocative stimuli. As short stature may affect psychological development, trials of GH in “slow” growing children have been initiated to determine whether GH treatment can increase final adult height in normal children without complications of treatment. GH treatment in these children is controversial. Long-term studies in short

children treated for up to ten years have shown conflicting results, some showing no significant difference in the SD score for final height [433] while others report final heights exceeding the predicted height [434]. Approximately onethird of all children on GH treatment in the USA are nonGH deficient with idiopathic short stature [435]. In view of the contradictory data on the benefical effect of GH therapy on final height, non-GH-deficient children and their families should have realistic expectations. In children with no identifiable cause of growth failure GH therapy is indicated if growth is retarded more than 2.5 SD below the mean for age and very slow growth velocity (less than the twenty-fifth percentile).

GH Neurosecretory Dysfunction (GHND) A subgroup of GH-deficient children has been described who have normal responses to GH provocative tests, low serum IGF-I levels and abnormal 24-hour GH secretory patterns [436]. This syndrome, termed “growth hormone neurosecretory dysfunction,” is characterized by height below the first percentile, bone age delayed by 2 or more years, and a growth velocity of 4 cm/year or less, in combination with the serum abnormalities described above. Classical GH deficiency, GHND, and normal children can be distinguished by assessing the number of GH pulses during a 24-hour period, mean peak GH pulse amplitude, highest nocturnal peak, and mean 24-hour GH concentration. Patients with GHND may secrete less GH than patients with classical GH deficiency, and the nocturnal GH surge is maintained but with a lower magnitude [436]. GHND has also been associated with Turner’s and Noonan’s syndrome, empty sella syndrome, and precocious puberty [437]. hGH was administered intramuscularly three times weekly for at least 6 months to children with GHND [436]. This treatment resulted in a doubling of growth velocity, comparable to that seen in children with GH deficiency receiving GH. Turner’s Syndrome

Patients with Turner’s syndrome manifest dysmorphic body features, ovarian failure, and reduced growth rate, starting during intrauterine life and continuing through childhood and puberty, resulting in reduced final adult height [438]. Earlier studies evaluating the use of hGH in Turner’s syndrome were inconclusive, possibly due to inadequate dosage regimens and short-term follow-up [439]. Recently, GH therapy in girls with Turner’s syndrome has been shown to significantly increase predicted height, with a greater increase in height in girls treated with combined GH and oxandrolone (an anabolic steroid) [440]. Turner’s syndrome is an approved FDA indication for GH use. Low Birth Weight

Children with low birth weight generally have reduced adult height [441]. An early study investigating the use of

Chapter 3

hGH in the treatment of short children with low birth weight yielded inconclusive results, probably due to the inadequate dosage regimen. Subsequent studies using higher, more frequent hGH dosages [442,443] have shown an increased growth rate for these children, but no increase in final height [444]. Chronic Diseases

hGH has been used in an attempt to increase height in children with short stature associated with a variety of chronic nonendocrine disorders. Growth retardation may be due to specific effects of the disease of may be secondary to poor nutrition or infection. Increased height velocity has been demonstrated after hGH treatment in prepubertal children and infants with chronic renal failure and following renal transplantation [445,446]. Chronic renal insufficiency prior

Table 3.5. deficiency

Causes of adult growth hormone (GH)

Presenting in childhood Congenital Idiopathic Genetic GHRH receptor defect Transcription factor defect Embryologic defects GH resistance Laron dwarfism Pygmy Neurosecretory defects Radiation for juvenile brain tumors, leukemia Head trauma Perinatal birth injury Child abuse Accidental Inflammatory diseases Viral encephalitis Meningitis, bacterial, fungal, tuberculosis Acquired in adulthood Postpituitary surgery Pituitary mass-adenoma, craniophyringioma Hematologic disorders Hemochromatosis Sickle cell disease Thalassemia Autoimmunity Lymphocytic hypophysitis Irradiation: central nervous system radiation for tumors Histiocytosis Granulomas Sarcoidosis Idiopathic Infection Tuberculosis Syphilis Vascular

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to transplantation is an FDA approved indication for GH therapy. Achondroplasia and Hypochondroplasia

Achondroplasia and hypochondroplasia result in short stature, with normal GH responses to provocative tests. GH therapy increases growth in both conditions, however there is worsening of body disproportion [447]. GH Treatment in Adults

As part of the normal aging process, adults experience loss of lean body mass, atrophy of skeletal muscle and organs, and increased adipose tissue [448]. Skeletal muscle atrophy is associated with a loss of muscle strength. These changes have been attributed to GH deficiency, as GH and IGF-I levels decline with age in healthy adults. With the increased availability of hGH, studies have been performed to evaluate the effects of rhGH on body composition in normal elderly individuals [449–451]. These studies have reported an increase in lean body mass, decreased adipose tissue content and increased retention of sodium, calcium, and nitrogen. None of these parameters translate into a measurable improvement in muscle size or strength. Adults may be truly GH-deficient due to isolated GH deficiency or to panhypopituitarism, from a number of causes (Table 3.5). GH-deficient adults have altered body composition with increased fat and decreased muscle volume and strength, lower psychosocial achievement, altered glucose and lipid metabolism, decreased bone mineral density and possibly increased mortality due to cardiovascular disease (Table 3.6). The same pharmacologic stimulating tests, used in children, have been used in adults. However, the definition of an adequate response differs. IGF-I, IGFBP-III, cholesterol and LDL levels can also be determined (Table 3.7). Mean 24-hour GH secretion sampled every 20 minutes, insulin-induced hypoglycemia and IGF-I levels have been compared to normals in the diagnosis of adult GHD [452]. IGF-I is a poor indicator of adult GHD as levels may overlap with normal. The diagnosis of adult GH deficiency is confirmed by provocative testing of GH secretion. Other hormonal deficits should be adequately replaced prior to GH provocative testing. The

Table 3.6. Physical findings in the adult growth hormone deficiency syndrome. Adapted from Cuneo et al. [451a] Truncal adiposity Waist/hip ratio increased Thin, dry, cool skin Exercise performance reduced Muscle strength reduced Reduced bone density Mood depressed Psychosocial impairment

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Table 3.7. Diagnosis of adult GH deficiency. Adapted from Cuneo et al. [451a] Stimulated GH <3 ng/mL Insulin Clonidine L-dopa Stimulated GH <9–16 ng/mL Arginine/GHRH IGF-I low or normal IGFBP-III low Hypercholesterolemia, with low HDL and often high LDL levels Reduced glomerular filtration rate Reduced lean body mass Reduced bone density

Table 3.8.

Table 3.9.

Complications of GH replacement therapy

Metabolic Glucose homeostasis Salt and fluid retention; peripheral edema Unmasking of thyroid dysfunction Joint Adults Children

Arthralgias and myalgias Carpal tunnel syndrome Slipped capital femoral epiphysis

Potential promotion of neoplastic growth Skin Pituitary adenoma Colonic polyps Other

GH therapy in adults Table 3.10.

Approved use GH Deficiency AIDS-associated muscle wasting Investigational use Frailty of aging Osteoporosis Catabolic states, cachexia Burns Postoperative recovery Wound healing Parenteral nutrition Ovulation induction Immune deficiency

insulin tolerance test is the current gold standard. Most normal subjects respond to insulin-induced hypoglycemia with a peak GH response of greater than 5 ng/mL. Severe GH deficiency is defined by a peak GH response to hypoglycemia of less than 3 ng/mL. Combined administration of arginine and GHRH is as sensitive as the insulin tolerance test. Childhood onset GH deficiency requires reconfirmation in adulthood by two GH provocative tests because growth hormone may no longer be deficient [453]. Adult patients with hypothalmic–pituitary disease and additional anterior pituitary hormone deficiencies require only one GH provocative test to confirm GH deficiency. Insulin induced hypoglycemia shows a clear separation of the peak GH response in GH-deficient adults compared to normals [454]. The scarcity of GH in the past has limited GH replacement therapy to GH-deficient children. The primary therapeutic goal has been to increase linear growth, and in the past treatment has been terminated after epiphyseal fusion occurs and a certain height has been achieved. However, GH secretion normally continues into adulthood, and GH influences many metabolic systems other than growth (Tables 3.8 and 3.9). The numerous beneficial effects of GH

Beneficial effects of GH replacement in adults

Body composition

Increased lean body mass Decreased fat mass Increased bone mass

Physical performance

Increased Max O2 uptake Increased max power

Metabolic effects

Increased IGF-I levels Increased BMR Decreased LDL with probable increased HDL Transient hyperglycemia Increased T3 levels Salt and water retention

Cardiovascular effects

Increased stroke volume Increased diastolic volume Increased LV wall mass Atherosclerosis impact

Psychological effects

Mood and energy uplift Enhanced vitality Improved physical mobility Social isolation improved

Adipose tissue effect

Decreased adipocyte size Increased lipolysis Decreased lipogenesis

replacement in adulthood have been previously described (Table 3.10). In addition, GH replacement in GH deficient adults is associated with increased energy levels, improved mood, vitality and emotional reactions and less feeling of social isolation [343]. GH replacement therapy is associated with significantly improved Nottingham Health Profile (NHP) energy scores (Fig. 3.17). GH doses in adulthood are adjusted to individual needs (Table 3.11). Elderly, male, or obese individuals require lower doses. Furthermore, the use of oral estrogen replacement effects the GH replacement dose. Premenopausal

Chapter 3

FIGURE 3.17. Effect of long term (10 years) treatment with GH on quality of life as measured by the Nottingham Health profile (Energy Level) in 21 GH deficient adults. (Note: higher scores indicate greater energy impairment). Reproduced with permission from Gibney et al. [343a].

Table 3.11. replacement

Factors determining side effects of GH

Patient age Enhanced IGF-I response Greater body weight and body mass index Adult onset vs. childhood onset of GH deficiency

women or postmenopausal women using transdermal estrogen replacement require lower doses of GH than postmenopausal women taking oral estrogen replacement [454]. Oral, but not transdermal estrogens antagonize the actions of GH and reduce IGF-I levels (Fig. 3.18). GH replacment therapy should be started at a low dose (0.15–0.30 mg/d) (GRS guidelines). GH is self-administered as a daily subcutaneous injection in the evening. Serum IGF-I levels and tolerability are used to guide dosing. IGF-I concentrations are monitored every 4 to 6 weeks, and GH doses are adjusted until levels reach mid-normal range for age and sex (Fig. 3.19). Side effects of GH replacement include edema and carpal tunnel syndrome due to fluid retention, which are usually transient, but may require dose reduction. Mild self-limiting arthralgias may also occur. As GH may reduce insulin sensitivity, glycemic control should be monitored. GH replacement therapy is contraindicated in patients with active malignancy, benign intracranial hypertension and proliferative diabetic retinopathy (Table 3.12). Changes in body composition, bone density and lipid profiles may only be observed after several months of therapy. These effects are best assessed by annual dual-energy x-ray absorptiometry
EXA and serum lipid level measurements. A baseline MRI is recommended prior to initial GH replacement. GH therapy does not impose a need for

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FIGURE 3.18. Time course of GH dose and serum IGF-I concentration in a representative patient (38-year-old woman) who was switched from oral to transdermal estrogen therapy during the course of GH replacement. Reproduced with permission from Cook et al. [454].

more frequent pituitary MRI’s than would be done for the regular clinical follow-up of residual pituitary tumor. GH Treatment in Catabolic States

The well-recognized anabolic actions of GH have tempted the use of GH in catabolic states such as surgery, trauma burns and organ failure. The negative nitrogen balance in critically ill patients is partly attributable to GH resistance as well as decreased production and action of IGF-I [455]. Postsurgical recovery is associated with hypercatabolism. GH administered to postsurgical patients as well as to normal subjects receiving hypocaloric intravenous alimentation [456], results in positive nitrogen balance [457]. Beneficial effects of GH in patients with extensive burns has been demonstrated in children [458] and adults [459]. Chronic high-dose glucocorticoid treatment is associated with protein catabolism. These effects in normal subjects may be reversed by simultaneous GH administration [459]. A potential complication of combined GH and glucocorticoid administration is resultant hyperglycemia. GH treatment improved nitrogen balance in two small studies of patients with chronic obstructive pulmonary disease [460,461], with improved respiratory muscle function only reported in one group [460]. Short-term studies of GH treatment in cancer patients suggested improvement in nitrogen balance, most pronounced in the least malnourished patients [461]. Although an uncontrolled study in critically ill patients suggested that GH treatment decreased mortality [462], a large European trial suggested increased mortality in critically ill patients treated with large doses of GH, and was terminated prematurely [463]. GH Treatment in Osteoporosis

Bone mass declines after the second decade of life in healthy men and women, with accelerated bone loss in post-

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FIGURE 3.19. Algorithm for the management of adult GH deficiency.

Table 3.12.

Contraindications to GH therapy in adults

Absolute Active neoplasm Intracranial hypertension Diabetic retinopathy Relative Uncontrolled diabetes Untreated thyroid dysfunction Cost benefit Need for injections

menopausal women. As declining GH secretion has been implicated in the pathogenesis of osteoporosis, GH has been administered to healthy men and women with idiopathic osteoporosis in an attempt to decrease bone loss [464]. Although GH therapy increased indices of bone formation and resorption [464] a small increase in only spine bone mineral density was observed in males, after 6 months [449], with an increase in spine mineral density in postmenopausal females when GH was administered with calcitonin [465]. A longer study in osteoporotic females showed increases in both spine and total hip bone mineral density [466], after 2 years of treatment with GH and calcitonin, although the response was less than with estrogen or biophosphonate therapy [467]. Limited efficacy, side effects and lack of comparative studies with beneficial therapies for osteoporosis, limit the use of GH in the treatment of osteoporosis.

[468], with an increase in extracellular water and lean body mass, a decrease in body fat and improved work output after 3 months [469]. Long term beneficial effects of GH on survival and quality of life have not been established yet. GH Therapy in Cardiac Failure

Beneficial effects of GH on cardiac function in GH deficiency led to its attempted use in congestive heart failure. GH administered for 3 months to a small group of patients with idiopathic dilated cardiomyopathy and moderate to severe congestive heart failure improved cardiac output [369] at rest and during exercise. A larger controlled study for a longer period of time did not confirm these findings [470]. Increased extracellular volume and edema, as well as the potential for cardiomyopathy as seen in acromegaly, must be considered in the use of GH in cardiac failure. GH and Wound Healing

GH treatment increases the strength and speed of collagen formation in incisional wounds in rats [471–473]. Fracture healing in rats is also facilitated by GH treatment [474]. No controlled data are yet available in human subjects. GH Treatment in Ovulation Induction

The growth hormone axis influences gonadal function [474] GH therapy reduces gonadotropin requirements in hypogonadotropic males and females [475,476]. Furthermore, improved implantation and pregnancy rates occur with the addition of GH to gonadotropin based protocols in women undergoing in vitro fertilization [477].

GH Treatment in AIDS-associated Wasting Syndrome

GH is FDA approved in adult patients with HIV associated cachexia. Short term GH administered to patients with HIV-associated cachexia results in positive nitrogen balance

GH use Competitive Sports

The ethical issues of GH abuse in sports have received much attention. GH has been used by many athletes to enhance

Chapter 3

muscle mass [478]. Whether this is accompanied by increased muscle strength is unknown. Continued pharmacologic use of GH by athletes would result in the adverse effects of acromegaly, which would decrease performance. COMPLICATIONS OF GH TREATMENT The complications of GH treatment have been described in GH-deficient patients treated with GH (see Table 3.9). Non-GH-deficient patients receiving GH can be likened to acromegalic patients, with increased GH levels. Adverse reactions, including arthralgias, myalgias, back ache, parasthesias, peripheral edema, carpel tunnel syndrome, headache, hypertension, and rhinitis are frequent, often transient or disappear with lowering of the GH dose and are more common in adult-onset than childhood-onset GH deficiency [479]. Although acromegalic patients have a potential increased risk of developing neoplasms of the gastrointestinal system, recent evidence is less compelling [480]. Colonic polyps occur more frequently, although their relationship to IGFI levels is not confirmed [481]. The true incidence of neoplasia in acromegalic patients is difficult to determine, although a potential link between GH and tumor development has been postulated in animal models. Elevated IGF-I levels in acromegaly may be responsible for cellular proliferation [482]. GH therapy in GH-deficient children has been implicated in the pathogenesis of leukemia. However, this was not confirmed in a worldwide survey [483]. Endogenous IGF-I concentrations have been epidemiologically linked to prostate cancer [484] and breast cancer risk [485]. ABNORMAL GH SECRETION GH hypersecretion, due to either excess GHRH (ectopic or eutopic) or to excess GH (ectopic or eutopic) secretion and GH deficiency (idiopathic or secondary) are documented fully in Chapter 10 (see also Table 3.13).

Table 3.13. acromegaly

Elevated GH levels not associated with

Associated decreased IGF levels Protein-calorie malnutrition Starvation Uncontrolled diabetes mellitus Anorexia nervosa GH insensitivity–Laron’s syndrome Decreased GH clearance Chronic renal failure Mucoviscidosis Physiologic (intermittent) Sleep Stress

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Decreased IGF-I Levels Protein-calorie Malnutrition, Starvation, Anorexia Nervosa

Short-term fasting in normal healthy people results in moderately elevated basal GH levels [486]. However, proteincalorie malnutrition is associated with low IGF-I levels and markedly elevated GH levels [487]. This may reflect an uncoupling of the IGF-I feedback regulation of GH secretion. Low levels of IGF-I in normal fasted volunteers were normalized on refeeding, and caloric intake normalized IGF-I levels to a greater degree than did protein intake [488]. In patients with anorexia nervosa, basal GH levels are also elevated [489]. Furthermore, GH levels rise paradoxically after a glucose load [490] and after TRH injection in these patients [491]. The abnormalities in anorexia nervosa are due not only to the weight loss, but to the concomitant psychiatric disturbance, which may influence hypothalamic– pituitary function. Diabetes Mellitus

Poorly controlled diabetes mellitus (both insulin dependent and noninsulin-dependent) is associated with elevated basal GH levels [90] and increased GH response to exercise [492] and to TRH [493]. Elevated GH levels return to normal with improved diabetic control after insulin administration [494]. IGF-I levels are low in children with poorly controlled insulin-dependent diabetes, including those entering puberty, suggesting insulin resistance. Antiinsulin actions of the increased GH may worsen the resistance. Addition of IGF-I to insulin therapy in adolescents with poorly controlled diabetes was postulated to decrease GH secretion, increase insulin sensitivity and decrease insulin requirements. Daily subcutaneous IGF-I injections for a month normalize IGF-I levels with less fluctuations in blood glucose levels, and reduced insulin doses [495]. Insulin resistance in non-insulin-dependent diabetes is associated with increased insulin secretion in an attempt to overcome putative post-receptor defects in insulin action [496]. IGF-I therapy was proposed to stimulate peripheral glucose utilization and reduce insulin resistance and secretion, and was shown to decrease fasting blood glucose by 30% with a 50% decrease in insulin secretion [497]. A high incidence of side effects with dosages used initially discouraged adjunctive IGF-I therapy [498]. Diabetes and malnutrition are both conditions of nutrient deprivation. In malnutrition, there is inadequate intake of nutrients, whereas diabetic patients are unable to utilize available nutrients appropriately. Malnutrition and diabetes both manifest retarded growth, despite elevated GH levels and decreased IGF-I generation. Laron Syndrome

Laron syndrome, an autosomal recessive disorder, is a condition of peripheral unresponsiveness to GH. Serum GH levels are normal or elevated, circulating IGF-I is absent, and

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there is no IGF-I response to exogenously administered GH [499]. Inactivating mutations of the GH receptor [500] cause insensitivity to exogenous GH. IGF-I therapy increased height velocity, with improved body composition as evidenced by loss of fat mass [501].

Decreased GH Clearance Chronic Renal Failure

Basal GH levels are high in patients with chronic renal failure [502], and GH rises paradoxically in response to a glucose load in these patients [502]. GH is degraded by the kidney. However, elevated basal GH levels imply some change in GH regulation; if the feedback loop were intact, GH levels should fall to normal. Serum IGF-I levels are reported to be elevated or variable in chronic renal failure [503–505]. As IGFs circulate bound to large carrier proteins, serum samples should be pretreated with acid-ethanol to ensure reliable quantitation of total IGFs [506]. Uremic serum may contain excess carrier protein not completely removed by acid-ethanol extraction, resulting in underestimation of immunoreactive IGF-I levels [505]. A more specific assay employs a second antibody to separate bound from free radioligand [507]. Despite high GH levels, children with renal failure are short. Following renal transplantation, return to normal growth is variable. Growth retardation can be inversely correlated with posttransplant creatinine clearance [508]. REFERENCES 1 Miller WL, Eberhardt NL. Structure and evaluation of growth hormone gene family. Endocrinol Rev 1983;4:97–130. 2 George DL, Phillips JA III, Francke U, Seeburg PH. The genes for growth hormone and chorionic somatomammotropin are on the long arm of human chromosome 17 in the region of q21-qter. Human Genet 1981;57:138–141. 3 Chen EY, Yu-Cheng L, Smith DH et al. The human growth hormone locus: nucleotide sequence, biology, and evolution. Genomics 1989;4:479–497. 4 Lewis UJ, Dunn JT, Bonewald LF et al. A naturally occurring structural variant of human growth hormone. J Biol Chem 1978;253:2679–2687. 5 Lewis UJ, Bonewald LF, Lewis VJ. The 20,000-dalton variant of human growth hormone: location of the amino acid deletions. Biochem Biophys Res Commun 1980;92:511–516. 6 DeNoto FM, Moore DD, Goodman HM. Human growth hormone DNA sequences and mRNA structure: possible alternative splicing. Nucleic Acids Res 1981;9:3719–3730. 7 Frankenne F, Rentier-Delrve F, Scippo ML et al. Expression of the growth hormone variant in human placenta. J Clin Endocrinol Metab 1987;64: 635–637. 8 Liebhaber SA, Urbanek M, Ray J et al. Characterization and histologic localization of human growth hormone—variant gene expression in the placenta. J Clin Invest 1989;83:1985–1991. 9 Frankeme F, Closset J, Gomez F et al. The physiology of growth hormone (GHs) in pregnant women and partial characterization of the placental GH variant. J Clin Endocrinol Metab 1988;66:1171–1180. 10 Cooke NE, Ray J, Watson MA et al. Human growth hormone gene and the highly homologous growth hormone variant gene display different splicing patterns. J Clin Invest 1988;82:270–275. 11 Ray J, Okamura H, Kelly PA et al. Human growth hormone-variant demonstrates a receptor binding profile distinct from that of normal pituitary growth hormone. J Biol Chem 1990;265:7939–7944. 12 MacLeod JN, Worsley I, Ray J et al. Human growth hormone-variant is a biologically active somatogen and lactogen. Endocrinology 1991;91:1290–1302.

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380 Hourd P, Edwards R. Current methods for the measurement of growth hormone in urine. Clin Endo 1994;40(2):155–170. 381 Albini CH, Quattrin T, Vandlen RL, McGillivray MH. Quantitation of urinary growth hormone in children with normal and subnormal growth. Pediatr Res 1988;23:89–92. 382 Hattori N, Shimatsu A, Yamanaka C et al. Nocturnal urinary growth hormone excretion in children with short stature. Acta Endocrinol (Copenh.) 1988;119:113–117. 383 Granada ML, Sanmarti A, Lucas A et al. Clinical usefulness of urinary growth hormone measurements in normal and short children according to different expressions of urinary growth hormone data. Pediatr Res 1992;32:73–76. 384 Rosenfeld RG, Albertsson-Wilkland K, Cassorl F et al. Diagnostic controversy: The diagnosis of childhood deficiency revisited. JCEM 1995;80:1532–1540. 385 Bercu BB, Shulman D, Root AW, Spiliotis BE. Growth hormone (GH) provocative testing frequently does not reflect endogenous GH secretion. J Clin Endocrinol Metab 1986;63:709–716. 386 Lunt H, Tucker AJ, Bullen H et al. Overnight urinary growth hormone measurement in the diagnosis of acromegaly. Clin Endocrinol 1990;33: 205–212. 387 Clemmons DR, Van Wyk JJ, Ridgway EC et al. Evaluation of acromegaly by radioimmunoassay of somatomedin-C. N Engl J Med 1979;301:1138–1142. 388 Barkan AL, Beitins IZ, Kelch RP. Plasma insulin-like growth factorI/Somatomedin-C in acromegaly: correlation with the degree of growth hormone hypersecretion. J Clin Endocrinol Metab 1988;67:69–73. 389 Melmed S, Jackson I, Kleinberg D et al. Current treatment guidelines for acromegaly. JCEM 1998;83:2646–2652. 390 Earll J, Sparks LL, Forsham PH. Glucose suppression of serum growth hormone in the diagnosis of acromegaly. JAMA 1967;201:628–630. 391 Beck P, Parker ML, Daughaday WH. Paradoxical hypersecretion of growth hormone in response to glucose. J Clin Endocrinol Metab 1966;26:463–469. 392 Phillips LS, Vassilopoulou-Sellin R. Somatomedins. Part I. N Engl J Med 1980;302:371–380. 393 Phillips LS, Vassilopoulou-Sellin R. Somatomedins. Part II. N Engl J Med 1980;302:438–446. 394 Dean HJ, Kellet JG, Bala RM et al. The effect of growth hormone treatment on somatomedin levels in growth hormone deficient children. J Clin Endocrinol Metab 1982;55:1167–1173. 395 Bala RM, Lopatka J, Leung A et al. Serum immunoreactive somatomedin-C levels in normal adults, pregnant women at term, children at various ages and children with constitutional delayed growth. J Clin Endocrinol Metab 1981;52:508–512. 396 Rubin KR, Lichtenfels JM, Ratzan SK et al. Relationship of somatomedin-C concentration to bone age in boys with constitutional delay of growth. Am J Dis Child 1986;19:975–980. 397 Reiter EO, Lovinger RD. The use of commercially available somatomedin-C radioimmunoassay in patients with disorders of growth. J Pediatr 1981;99: 720–724. 398 Blum WF, Ranke MB, Kietzmann K et al. A specific radioimmunoassay for growth hormone (GH)-dependent somatomedin-binding protein: its use for diagnosis of GH deficiency. J Clin Endocrinol Metab 1990;70:1292–1298. 399 Hasegawa Y, Hasegawa T, Aso T et al. Usefulness and limitation of measurement of insulin-like growth factor binding protein 3 for diagnosis of GH deficiency. Endocrinol Jpn 1992;39:585–591. 400 Hasegawa Y, Hasegawa T, Kotch S, Tsuchiya Y. Reproducibility of GH stimulation tests (arginine and insulin) IGF-1 and IGFBP-3 measurements. Clin Pediatr Endocrinol 2 1993;(Suppl):75–78. 401 Growth Hormone Research Society. Consensus guidelines for the diagnosis and treatment of adults with growth hormone deficiency: summary statement of the Growth Hormone Research Society workshop on Adult Growth Hormone Deficiency. J Clin Endocrinol Metab 1998;83:379–381. 402 Gil-Ad I, Topper E, Laron Z. Oral clonidine as a growth hormone stimulation test. Lancet 1979;2:278–279. 403 Gil-Ad I, Gurewitz R, Marcovici O et al. Effects of aging on plasma GH response to aging. Mech Aging Dec 1984;27:97–100. 404 Fass B, Lippe BM, Kaplan SA. Relative usefulness of three growth hormone stimulation screening tests. Am J Dis Child 1979;133:931–933. 405 Ghigo E, Bellone J, Mazza E et al. Arginine potentiates the GHRH but not the pyridostigmine-induced GH secretion in normal short children. Further evidence for a somatostatin suppressing effect of arginine. Clin Endocrinol (Oxf.) 1990;32:763–767. 406 Aimaretti G, Corneli G, Razzore P et al. Comparison between insulininduced hypoglycemia and GHRH plus Arginine as provocative tests for the diagnosis of GH deficiency in adults. JCEM 1998;83:1615–1619.

407 Popovic V, Leal A, Dragan M et al. Lancet 2000;356:1137–1142. 408 Ho KKY. Diagnosis of adult GH deficiency. Lancet 2000;356:1125–1126. 409 Penny R, Blizzard RM, Davis WT. Sequential arginine and insulin tolerance tests on the same day. J Clin Endocrinol Metab 1969;29:1499–1501. 410 Reiter EO, Morris AH, Biggs DE. Modulation of GHRH-induced growth hormone release by an alpha-adrenergic agonist and hypoglycemia. J Pediatr Endocrinol 1988;3:21–25. 411 Shibasaki T, Hotta M, Masuda A et al. Plasma GH response to GHRH and insulin-induced hypoglycemia in man. J Clin Endocrinol Metab 1985;60: 1265–1267. 412 Woolf PD, Lantigua R, Lee LA. Dopamine inhibition of stimulated growth hormone secretion: evidence for dopaminergic modulation of insulin and L-dopa-induced growth hormone secretion in man. J Clin Endocrinol Metab 1979;49:326–330. 413 Weldon VV, Gupta SK, Klingensmith G et al. Evaluation of growth hormone release in children using arginine and L-dopa in combination. J Pediatr 1975;87:540–544. 414 Cacciari E, Tassoni P, Cigognani A et al. Value and limits of pharmacological and physiological test to diagnose GH deficiency and predict therapy response: first and second retesting during replacement therapy of patients defined as GH deficient. JCEM 79:1663–1669. 415 Sperling MA, Kenny FM, Drash AL. Arginine-induced growth hormone responses in children: effect of age and puberty. J Pediatr 1970;77: 462–465. 416 Illig R, Bucher H. Testosterone priming of growth hormone release. In: Laron Z, Butenandt O, eds. Evaluation of Growth Hormone Secretion. Basel: Karger, 1983:75–83. 417 MacGillivray MH, Aceto T Jr, Frohman LA. Plasma growth hormone responses and growth retardation of hypothyroidism. Am J Dis Child 1968;115:273–276. 418 Tauber M, Moulin P, Pierkowski C et al. GH retesting and auxiological data in 131 GH deficient patients after completion of treatment. JCEM 1997;82: 352–356. 419 Granada ML, Sanmarti A, Lucas A et al. Assay-dependent results of immunoassayable spontaneous 24 hr GH secretion in short children. Acta Ped Scand 1990;370(suppl):63–70. 420 Baumann G, Shaw MA, Amburn K et al. Heterogeneity of circulating growth hormone. Nucl Med Biol 1994;21:369–379. 421 Reiter EO, Morris AH, MacGillivray MH, Weber D. Variable estimates of serum GH concentrations by different radioassay systems. JCEM 1988;66: 68–71. 422 Orskov H, Weeke J, Frystyk J, Kaal A, Nielsen S et al. Growth hormone determination in serum from patients with growth disorders; in Juul A, Jorgenson JOL eds. Growth Hormone in Adults: Physiological and Clinical Aspects. Cambridge: Cambridge University Press, 1996:109–121. 423 Bristow AF, Gaines Das R, Jeffcoate SL, Schulster D. The first international standard for somatotropin: Report of an international collaborative study. Growth Regul 1995;5:133–141. 424 Fisker S, Orskov H. Factors modifying GH estimates in Immunoassays. Horm Res 1996;46:183–187. 425 Strasburger CJ, Dattani MT. New growth hormone assays: potential benefits. Acta Pediatric 1997;423(Suppl):5–11. 426 Goeddel DV, Heyneker HL, Hozumi T et al. Direct expression in Escherichia coli of a DNA sequence coding for human growth hormone. Nature 1979;281:544–548. 427 Hintz RL, Rosenfeld RG, Wilson DM et al. Biosynthetic methionyl human growth hormone is biologically active in adult man. Lancet 1982;i:1276. 428 Vance ML, Mauras N. Drug therapy: growth hormone therapy in adults and children. N Engl J Med 1999;341:1206–1216. 429 Guidelines for the use of growth hormone in children with short stature; a report by the Drug and Therapeutics Committee of the Lawson Wilkins Pediatric Endocrine Society. L Pediatr 1995;127:857–867. 430 Joss E, Zuppinger K, Schwartz HP, Roten H. Final height of patients with pituitary growth failure and changes in growth variables after long-term hormonal therapy. Pediatr Res 1983;17:676–679. 431 Maes M, Lindberg A, Price DA et al. Long term growth response to growth hormone therapy in prepubertal children with idiopathic growth hormone deficiencies: analysis of Kabi International growth study. Kabi International Growth Study Biannual Report No 11. Mannheim, Germany: J and J Verlag, 1994:15–26. 432 Preece MA, Tanner JM, Whitehouse RH, Cameron N. Dose dependence of growth response to human growth hormone in growth hormone deficiency. J Clin Endocrinol Metab 1976;42:477–483.

Chapter 3 433 Loche S, Cambiaso P, Setzu S et al. Final height after growth hormone therapy in non-growth hormone deficient children with short stature. J Pediatr 1994;125:196–200. 434 Hintz RL, Attie KM, Baptista J, Roche A. Effect of growth hormone treatment on adult height of children with idiopathic short stature. N Engl J Med 1999;340:502–507. 435 Cuttler L, Silvers JB, Sing J et al. Short stature and growth hormone therapy: a national study of physician recommendation patterns. JAMA 1996; 276:531–537. 436 Spiliotis BE, August GP, Hung W et al. Growth hormone neurosecretory dysfunction: a treatable cause of short stature. JAMA 1984;251:2223–2230. 437 Bercu BB, Diamond F. Growth hormone neurosecretory dysfunction. J Clin Endocrinol Metab 1986;15:537–590. 438 Lyon AJ, Preece MA, Grant DB. Growth curve for girls with Turner syndrome. Arch Dis Child 1985;60:932–935. 439 Stahnke N. hGH treatment in short children without growth hormone deficiency. N Engl J Med 1984;310:925–926. 440 Rosenfeld RG, Attie KM, Frane J et al. Growth hormone therapy for Turner’s syndrome: beneficial effect on adult height. J Pediat 1998;132:319–324. 441 Davies PSW, Valley R, Preece MA. Adolescent growth and pubertal progression in the Silver-Russel syndrome. Arch Dis Child 1988;63: 130–135. 442 Lanes R, Plotnick LP, Lee PA. Sustained effect of human growth hormone therapy on children with intrauterine growth retardation. Pediatrics 1979;63:731–735. 443 Stanhope R, Ackland F, Hamill G et al. Physiological growth hormone secretion and response to growth hormone treatment in children with short stature and intrauterine growth retardation. Acta Paediatr Scand 1989;349(suppl.):47–52. 444 de Zegher F, Maes M, Gargosky SE et al. High dose growth hormone treatment of short children born small for gestational age. JCEM 1996;81:1887–1892. 445 Hokken-Koelega AC, Stijnen T, de Jong RC et al. A placebo-controlled, double-blind trial of growth hormone treatment in prepubertal children after renal transplant. Kidney Int Suppl 1996;53:S128–S134. 446 Fine RN. Recombinant human growth hormone in children with chronic renal insufficiency-clinical update: 1995. Kidney Int Suppl 1996;53: S115–S118. 447 Bridges NA, Brook CGD. Growth hormone in skeletal dysplasia. Horm Res 1994;42:231–234. 448 Rudman D. Growth hormone, body composition and aging. J Am Geriatr Soc 1985;33:800–807. 449 Rudman D, Feller AG, Nagraj HS et al. Effects of human growth hormone in men over 60 years old. N Engl J Med 1990;323:1–6. 450 Binnerts A, Wilson JHP, Lamberts SWJ. The effects of human growth hormone administration in elderly adults with recent weight loss. J Clin Endocrinol Metab 1988;67:1312–1316. 451 Marcus R, Butterfield G, Holloway L et al. Effects of short-term administration of recombinant human growth hormone to elderly people. J Clin Endocrinol Metab 1990;70:519–527. 451a Cuneo RC, Salomon F, Macgauley GA, Sonksen PH. The growth of hormone deficiency syndrome in adults. Clin Endocrinol 1992;37:387–397. 452 Hoffman DM, O’Sullivan AJ, Baxter RC, Ho KKY. Diagnosis of GH deficiency in adults. Lancet 1994;343:1064–1068. 453 Nicolson A, Toogood AA, Rahim A, Shalet SM. The prevalence of severe growth hormone deficiency in adults who received growth hormone replacement in childhood. Lancet 1996;44:311–316. 454 Cook DM, Ludlam WH, Cook MB. Route of estrogen administration helps to determine GH replacement dose in GH deficient adults. J Clin Endocrinol Metab 1999;84:3956–3960. 455 Jenkins RC, Ross RJM. Growth hormone therapy for protein catabolism. QJ Med 1996;89:813–819. 456 Manson JM, Wilmore DW. Positive nitrogen balance with human growth hormone and hypocaloric intravenous feeding. Surgery 1986;100:188–197. 457 Ponting GA, Halliday D, Teale JD, Sim AJW. Postoperative positive nitrogen balance with intravenous hyponutrition and growth hormone. Lancet 1988;i:438–440. 458 Herndon DN, Barrow RE, Kunkel KR et al. Effects of recombinant human growth hormone on donor-site healing in severely burned children. Ann Surg 1990;212:424–429. 459 Knox JB, Demling RH, Wilmore DW et al. Increased survival after major thermal injury: the effect of GH therapy in adults. J Trauma 1995;39:526–530.

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460 Pape GS, Friedman M, Underwood LE et al. The effect of GH on weight gain and pulmonary function in patients with chronic obstructive lung disease. Chest 1991;99:1495–1500. 461 Tayek JA, Brasel JA. Failure of anabolism in malnourished cancer patients receiving growth hormone: a clinical research center study. J Clin Endocrinol Metab 1995;80:2082–2087. 462 Knox JB, Wilmore DW, Demling RH et al. Use of GH for postoperative respiratory failure. Ann J Surg 1996;171:576–580. 463 Takada J, Ruokonen E, Webster N et al. Increased mortality associated with growth hormone treatment in critically ill adults. N Engl J Med 1999;341:785–792. 464 Johansson AG, Lindh E, Blum WF et al. Effects of growth hormone and IGF in men with idiopathic osteoporosis. J Clin Endocrinol Metab 1996;81:44 –48. 465 Gonnelli S, Cepollaro C, Montomoli M et al. Treatment of postmenopausal osteoporosis with recombinant human growth hormone and salmon calcitonin: a placebo controlled study. Clin Endocrinol 1997;46:55–61. 466 Holloway L, Kohlmeier L, Kent K et al. Skeletal effects of cyclic recombinant human growth hormone and salmon calcitonin in osteoporotic postmenopausal women. J Clin Endocrinol Metab 1997;82:1111–1117. 467 Deal CL. Osteoporosis: prevention, diagnosis and management. Am J Med 1997;102:355–395. 468 Mulligan K, Grunfeld C, Hellerstein MK et al. Anabolic effects of recombinant human growth hormone in patients with wasting associated with HIV infection. J Clin Endocrinol Metab 1993;77:956–962. 469 Schambelan M, Mulligan K, Grunfield C et al. A controlled trial of recombinant human growth hormone in patients with HIV associated growth hormone wasting. Ann Inter Med 1996;125:873–882. 470 Osterziel KJ, Strohm O, Schuler J et al. Randomized, double-blinded, placebo-controlled trial of human recombinant growth hormone in patients with chronic heart failure due to dialated cardiomyopathy. Lancet 1998;351:1233–1237. 471 Hollander DM, Devereux DF, Marafino BJ, Hoppe H. Increased wound breaking strength in rats following treatment with synthetic human growth hormone. Surg Forum 1984;35:612–614. 472 Jorgensen PH, Andreassen TT, A dose-response study of the effects of biosynthetic human growth hormone on formation and strength of granulation tissue. Endocrinology 1987;121:1637–1641. 473 Belcher HJCR, Ellis H. Somatotropin and wound healing. J Clin Endocrinol Metab 1990;70:939–943. 474 Homburg R, Levy T, Ben-Rafael Z. Adjuvant growth hormone for induction of ovulation with GnRH agonist and gonadotropins in polycystic ovary syndrome: a randomized, doubleblind, placebo controlled trial. Hum reprod 1995;10:2550–2553. 475 Homburg R, Eshel A, Abdalla HI et al. Growth hormone facilitates ovulation induction by gonadotropins. Clin Endocrinol 1988;29:113–117. 476 Shoham Z, Conway GS, Ostergard H et al. Cotreatment with growth hormone for induction of spermatogenesis in patients with hypogonadotropic hypogonadism. Fertil Steril 1992;57:1044–1051. 477 Bergh C, Hillensjo T, Wikland M et al. Adjuvant growth hormone treatment during in vitro fertilization: a randomized, placebo controlled study. Fert Steril 1994;62:113–120. 478 Crist DM, Peake GT, Egan PA, Waters DL. Body composition response to exogenous GH during training in highly conditioned adults. J Appl Physiol 1988;65:579–584. 479 DeBoer H, Blok GJ, Van Der Veen EA. Clinical aspects of growth hormone deficiency in adults. Endo Rev 1995;16:63–86. 480 Melmed S. Acromegaly and cancer: not a problem? J Clin Endocrinol Metab 2000;86:2929–2934. 481 Shalet SM, Renehan AG, Painter JE et al. Circulating insulin-like growth factor II and colorectal adenomas. JCEM 2000;85(9):3402–3408. 482 Besser M, Jenkins PJ, Fairclough PD et al. Acromegalic, colonic polyps and carcinoma. Clinica Endocrinol 1997;47(1):17–22. 483 Daughaday WH. The possible autocrine/paracrine and endocrine roles of insulin-like growth factors of human tumors. Endocrinology 1990;127:1–4. 484 Stahnke N, Zeisel HJ. Growth hormone therapy and leukemia. Eur J Pediatr 1989;148:591–596. 485 Chan JM, Stampfer MJ, Giovannucci E et al. Circulating insulinlike growth factor I and prostate cancer risks a prospective study. Science 1998;279: 563–566. 486 Hankinson S, Pollak M, Michaud D et al. A prospective assessment of plasma insulin-like growth factor levels and breast cancer risk. Am J Epidemiol 1997;145:s72. 487 Cahill GR, Herrera MG, Morgan AP et al. Hormone-fuel interrelationships during fasting. J Clin Invest 1966;45:1751–1769.

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488 Grant DB, Hambley J, Becker D, Pimstone BL. Reduced sulfation factor in undernourished children. Arch Dis Child 1973;48:596–600. 489 Isley WL, Underwood LE, Clemmons DR. Dietary components that regulate serum somatomedin-C concentrations in humans. J Clin Invest 1983;71: 175-182. 490 Brown GM, Garfinkel PE, Jeuniewic N et al. Endocrine profiles in anorexia nervosa. In: Vigersky RA, ed. Anorexia Nervosa. New York: Raven Press, 1977:123–125. 491 Casper RC, Davis JM, Pandey GN. The effect of the nutritional status and weight changes on hypothalamic function tests in anorexia nervosa. In: Vigersky RA, ed. Anorexia Nervosa. New York: Raven Press, 1977:137–147. 492 Macaron C, Wilber JF, Green O, Freinkel N. Studies of growth hormone (GH), thyrotropin (TSH) and prolactin (PRL) secretion in anorexia nervosa. Psychoneuroendocrinology 1978;3:181–185. 493 Hansen AP. Abnormal serum growth hormone response to exercise in juvenile diabetics. J Clin Invest 1971;49;1467–1478. 494 Dasmahapatra A, Urdanivia E, Cohen MP. Growth hormone response to thyrotropin-releasing hormone in diabetes. J Clin Endocrinol Metab 1981;52:859–862. 495 Hansen AP, Johansen K. Diurnal patterns of blood glucose, serum free fatty acids insulin, glucagon and growth hormone in normal and juvenile diabetics. Diabetologia 1970;6:27–33. 496 Bach MA, Chin E, Bondy CA. The effects of subcutaneous insulin-like growth factor-I infusion in insulin-dependent diabetes mellitus. J Clin Endocrinol Metab 1994;79:1040–1045. 497 Moller DE, Flier JS. Insulin resistance-mechanisms, syndromes and implications. NEJM 1991;325:938–948. 498 Zenobi PD, Jaeggi-Groisman SE, Riesen WF et al. Insulin-like growth factor-I improves glucose and lipid metabolism in type 2 diabetes mellitus. J Clin Invest 1992;90:2234–2241. 499 Jabri N, Schalch DS, Schwartz SL et al. Adverse effects of recombinant human insulin-like growth factor I in obese insulin-resistant type II diabetic patients. Diabetes 1994;43:369–374.

500 Laron Z, Pertzelan A, Karp M et al. Administration of growth hormone to patients with familial dwarfism with high plasma immunoreactive growth hormone measurement of sulfation factor, metabolic and linear growth response. J Clin Endocrinol Metab 1971;33:332–342. 501 Godowski PJ, Leung DW, Meacham LR et al. Characterization of the human growth hormone receptor gene and demonstration of a partial gene deletion in two patients with Laron-type dwarfism. Proc Natl Acad Sci USA 1989;86:8083–8087. 502 Laron Z, Anin S, Klipper-Aurbach Y, Klinger B. Effects of insulin-like growth factor on linear growth, head circumference, and body fat in patients with Laron-type dwarfism. Lancet 1992;339:1258–1261. 503 Samaan NA, Freeman RM. Growth hormone levels in severe renal failure. Metabolism 1970;19:102–113. 504 Andress DL, Pandian MR, Endres DB, Kopp JB. Plasma insulin-like growth factors and bone formation in uremic hyperparathyroidism. Kidney Int 1989;36:471–477. 505 Goldberg A, Trivedi B, Delmez J et al. Uremia reduces insulin-like growth factor-I, increases insulin-like growth factor-II and modifies their serum protein binding. J Clin Endocrinol Metab 1982;55: 1040–1045. 506 Powell DR, Rosenfeld RG, Baker BK et al. Serum somatomedin levels in adults with chronic reneal failure: the importance of measuring insulin-like growth factor-I (IGF-I) and IGF-II in acid-chromatographed uremic serum. J Clin Endocrinol Metab 1986;63:1186–1192. 507 Daughaday WH, Mariz IK, Blethen SL. Inhibition of access of somatomedin to membrane receptor and immunobinding sites: comparison of radioreceptor and radioimmunoassay of somatomedin in native and acid-ethanol-extracted serum. J Clin Endocrinol Metab 1980;51:781–788. 508 Daughaday WH, Kapadia M, Mariz I. Serum somatomedin binding proteins: physiologic significance and interference of radioligand assay. J Lab Clin Med 1987;109:355–363. 509 Saenger P, Wiedermann E, Schwartz E et al. Somatomedin and growth after renal transplantation. Pediatr Res 1974;8:163–169.

C h a p t e r

4 Prolactin Mark E. Molitch

HISTORICAL OVERVIEW In the late 1920s and early 1930s, a number of groups found that pituitary extracts could induce milk secretion [1–3]. Riddle and coworkers found that this substance, which they named prolactin (PRL), could be differentiated from the known growth and gonad stimulating substances [3–5]. In these experiments, they showed that PRL stimulated milk production by guinea pig mammary glands and a milk-like substance from the crop sacs of pigeons and doves [3–6]. The pigeon crop sac assay for PRL [3–6] then became the standard assay procedure for PRL and was used in many experiments over the next 30 years. Over the ensuing years, PRL was characterized, sequenced and specific radioimmunoassays (RIAs) developed for PRL from a number of species [7–10]. Because of the high lactogenic activity of even very highly purified preparations of human growth hormone (GH) [11,12], however, it was impossible to separate human PRL from GH using the relatively crude pigeon crop assay and the evaluation of PRL physiology in humans proceeded slowly. Even the existence of a PRL separate from GH in humans was questioned [12]. Several experiments of nature provided strong evidence, however, that even in humans these two hormones were separate. For example, it was observed that most patients with pituitary tumors in whom galactorrhea and amenorrhea were the cardinal clinical features did not have acromegalic features [13] and patients who were known to have isolated, congenital GH deficiency were able to undergo postpartum lactation [14]. Finally, in 1970, Frantz and Kleinberg developed a sensitive in vitro bioassay which involved staining milk produced by cultured, lactating mouse mammary tissue in response to PRL that was capable of measuring ovine PRL levels as low as 5 ng/ml. In this

assay they added excess antibody to GH to neutralize any potential lactogenic effects it had and, for the first time, were able to demonstrate measurable PRL levels in women with puerperal and nonpuerperal galactorrhea but not in most normal men and women [15]. Shortly thereafter, still using the pigeon crop assay, Lewis et al. reported considerable progress in the purification of human PRL and its separation from GH via several sequential separation techniques, leading the way to the eventual development of heterologous [16] and homologous [17] RIAs for human PRL which could finally measure PRL levels in the sera of normal individuals. The RIA for human PRL finally permitted the eventual sequencing of human PRL [18] and determination of its cDNA sequence [19]. With the development of PRL RIAs, studies of PRL physiology and pathophysiology in animals and humans have progressed at an astonishing pace over the past 20 years.

CELL OF ORIGIN

Lactotroph Morphology PRL is made by the pituitary lactotrophs (also known as mammotrophs). In the normal human pituitary, the lactotrophs comprise about 15% to 25% of the total number of cells, are similar in number in both sexes and do not change significantly with age [20–22]. During pregnancy and subsequent lactation, however, considerable lactotroph hyperplasia may be found [23,24], presumably due to the stimulatory effect of the hormonal milieu of pregnancy (see below for a discussion of the effects of estrogen on PRL secretion). The hyperplastic process involutes within several months after delivery, although breast-feeding retards this process [24]. This stimulatory effect of pregnancy on the lactotrophs also holds true for prolactinomas and very 119

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significant pregnancy-induced tumor enlargement may occur (see Chapter 12, p. 483). The lactotrophs are of two types: (i) large, polyhedral elongated cells distributed throughout the gland but generally near capillaries; and (ii) small-to-medium-sized, angular or elongated cells appearing in clusters on the border between the lateral wings and the median wedge and in the posterolateral edges of the gland [25]. The larger cells correspond to the densely granulated (granules 250–800 nm in diameter) cells seen on electron microscopy that are thought to be in a resting, storing stage while the smaller cells correspond to the sparsely granulated (granules 200– 350 nm in diameter) cells seen on electron microscopy that are actively secreting hormone [22,25]. Within these cells, PRL synthesis on the ribosomes of the rough endoplasmic reticulum, packaging into secretory granules within the Golgi, and release into the perivascular space by exocytosis appears to proceed in a normal fashion similar to that of other secretory proteins [25,26], although exocytosis may also occur at the lateral cell membranes, a phenomenon referred to as “misplaced exocytosis” [25]. Although it is possible that the densely and sparsely granulated lactotrophs represent different cell types, it is more likely that they represent the same cells in different phases of storage and secretion [22]. Most PRL is secreted by lactotrophs in the normal individual; however, there is a small proportion that is secreted by mammosomatotrophs, that are also capable of secreting GH [20,22,27]. In these cells some secretory granules contain GH, some PRL and some both hormones [27]. The ability of individual adenohypophyseal cells to secrete PRL in culture can be assessed using the reverse hemolytic plaque assay (Fig. 4.1) [28]. In this assay, the PRL secreted by individual cells is ascertained by identifying zones of hemolysis (plaques) around individual lactotrophs when

incubated in a monolayer with staphylococcal protein-Acoated ovine erythrocytes in the presence of PRL antiserum and complement [28]. Initial experiments with this system revealed that stimulatory agents such as thyrotropin releasing hormone (TRH) could cause an increase in plaque number and size, and inhibitory agents such as dopamine (DA) could cause a decrease in plaque number and size [28]. Subsequent studies have shown that there is considerable heterogeneity in the responsiveness of lactotrophs to such stimuli. A subpopulation of lactotrophs, comprising about 65% of cells, forms large plaques and is readily suppressible by DA; the remaining cells form small plaques and are not suppressible by DA [29]. In other studies, only one-third of lactotrophs were shown to respond to TRH [30]. Further analysis revealed that cells taken from the peripheral rim of the pituitaries of lactating rats were greatly responsive to TRH but poorly responsive to DA, whereas cells from the central region of the pituitary were little influenced by TRH but highly suppressible by DA [31]. The reverse hemolytic plaque assay, applied in a sequential fashion to PRL and GH, reveals that in normal rats of the pituitary cells secreting both GH and PRL, about one-third secrete GH alone, one-third secrete PRL alone and onethird secrete both hormones (i.e. mammosomatotrophs) [32], although mammosomatotrophs were found in only 5% of cells in another study [33]. In neonatal rats, the proportion of these cells that secrete only PRL was only 1.7%, suggesting that PRL-secreting cells arise from GH-secreting cells, i.e. mammosomatotrophs [34]. Tissue obtained near the periphery contained a higher proportion of cells that produced both PRL and growth hormone (GH) than did tissue obtained from the central region of the pituitary [31]. Estradiol treatment causes some cells that had only secreted GH to also secrete PRL, i.e. they converted from somatotroph

FIGURE 4.1. Reverse hemolytic plaque assay. In this assay, the prolactin secreted by individual cells is ascertained by identifying these zones of hemolysis (plaques) when incubated in a monolayer with staphylococcal protein A-coated ovine erythrocytes in the presence of PRL antiserum and complement. From Neill and Frawley [28]

Chapter 4

to mammosomatotroph cells [35]. Added GH releasing hormone (GHRH) and gonadotropin releasing hormone (GnRH) caused an increase in the proportion of PRL secretors and a decrease in the proportion of GH secretors, but corticotropin releasing hormone (CRH) increased the proportion of PRL secretors and did not affect GH secretors [36]. As yet, few studies have been performed with the reverse hemolytic plaque assay with human pituitary tissue. In one study of human fetal pituitary tissue (gestational age 18 to 22 weeks), 8% of cells formed PRL plaques [37], a smaller number than would expected from studies of adult pituitaries (see above). Of cells secreting either GH or PRL, 70% secreted GH only, 9% secreted PRL only and 21% secreted both hormones [37]. If we assume that heterogeneity of basal secretion, the response to various stimulatory and suppressive substances, and the ability of the cell population to shift between GH, PRL and GH plus PRL secretion all also take place in the adult human, such findings have obvious importance with regard to the pathogenesis of prolactinomas. For example, such tumors may arise from cells that are minimally or not responsive to endogenous inhibition by DA and thus may explain the variability of suppression of PRL by tumors in response to dopaminergic agents such as bromocriptine.

Lactotroph Ontogeny Continuous hypothalamic–pituitary interaction takes place during embryologic development. During the formation of Rathke’s pouch, the primordium of the anterior pituitary, the ectodermal primordial cells of the anterior and intermediate lobes of the pituitary make contact with the neuroectoderm of the floor of the diencephalon and experimental studies have shown inductive interactions between these tissues that are necessary for their subsequent interdependent development [38]. Elegant studies using a series of targeted mutations have shown that there is a whole host of homeodomain transcription factors that are sequentially expressed in the developing hypothalamus and pituitary that lead to the final determination of the five mature pituitary cell types and the functional integration of the hypothalamic–pituitary system [38,39]. In the earliest phase, in which Rathke’s pouch first appears, the transcription factors Six-3 and Hesx1 (also known as Rpx) are necessary for the initial development of Rathke’s pouch and other midline brain structures [38,39]. In humans, mutations in the genes for these factors result in defects in the development of midline structures and failure of normal hypothalamic and pituitary development with clinical hypopituitarism [40,41]. The LIM homeobox gene Lhx3 begins to be expressed in Rathke’s pouch at the time of organ commitment, in part stimulated by bone morphogenic protein 4 (BMP4) that is expressed in the ventral diencephalon [39] and inactivation/deletion of both the BMP4 and Lhx3 genes in mice causes a failure of the pitu-

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itary to develop, with the exception of the corticotroph lineage, despite the formation of Rathke’s pouch [39,42]. Lhx3 can synergize with Pit-1 in the transcriptional activation of the gene that encodes PRL and is required for the eventual proliferation and determination of the lactotroph, thyrotroph, gonadotroph, and somatotroph lineages [43]. Another homeobox-containing transcription factor gene, PTX1/P-OTX, is expressed in Rathke’s pouch early but later is restricted to corticotrophs and its role in the development of lactotrophs is still unclear [38]. A second BMP, BMP2, is transiently expressed in Rathke’s pouch and its temporal rise and fall both seem to be necessary for terminal pituitary cell differentiation including specification of the Pit-1 lineage [39]. Fibroblast growth factor 8 (FGF8) and Wnt4 have also been determined as important in the proliferation of the various pituitary cell types [39]. The POU homeodomain transcription factor Pit-1 gene (see below) becomes activated close to the same time as the Lhx3 gene and is also necessary for the activation of the PRL, GH, GHRH receptor, and TSHb genes as well as being necessary for the differentiation and proliferation of these cell lineages. Pit-1 also activates the c-fos promoter and Fos is known to be involved in cell cycle initiation [44]. A point mutation in the POU homeodomain of Pit-1 has been found to be the cause of the GH, PRL, and TSH deficiencies found in the Snell dwarf mouse, with absence of somatotroph, lactotroph, and thyrotroph cells [45]. Similar mutations in the POU-specific domain [46] and the POU homeodomain [47] of Pit-1 have now been found to cause a similar deficiency of GH, PRL and TSH in humans. A second paired-like homeodomain factor, known as Prophet of Pit-1 (Prop-1) has also been found to be necessary for the expression of Pit-1 and mutations of the Prop-1 gene causes the dwarfism in mice known as the Ames mouse (defects of somatotrophs, lactotrophs and thyrotrophs) [48] and similar mutations in humans cause variable deficiencies of GH, PRL, TSH, LH and FSH [49–51]. Although the glycoprotein a-subunit is the first hormone transcript to be expressed during the development of the pituitary, such cells are not the precursor of subsequent lactotrophs, as found by targeted ablation of a-subunit expressing cells [52]. Most lactotrophs arise from cells that at least at some point expressed the GH gene [53]. Subsequent lactotroph proliferation occurs once estrogen receptors appear [53]. Estrogen stimulates PRL gene transcription (see below) only if Pit-1 is bound to the PRL promoter [54]. Thus, the division of corticotroph from other cell premordia occurs very early. Stimulation by Prop-1 is necessary for the subsequent development of all noncorticotroph cells and Pit-1 is then necessary for the development and proliferation of thyrotrophs and somatomammotrophs, but the separation of these two cell lines occurs before the appearance of the ability to synthesize a-subunits. The final differentiation of somatomammotrophs occurs at least in part in relationship to estrogen stimulation.

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PROLACTIN GENE STRUCTURE

Chromosome Location The PRL gene in the human is located on chromosome 6, whereas those of the related hormones GH and human placental lactogen (HPL, also known as human chorionic somatomammotropin or HCS) are located on chromosome 17 [55,56]. In the rat, PRL similarly is dispersed to a separate chromosome from its relatives, PRL being located on chromosome 17 and GH on chromosome 10 [57]. These three genes (and possibly others, see below) are thought to have derived by gene reduplication from a common ancestral gene which diverged about 400 million years ago [19,58,59]. In both the human and the rat, the PRL genes and amino acid sequences show less homology with GH than placental lactogen does, implying an earlier divergence of PRL and GH and a more recent divergence of GH and PL [59].

PRL Gene Structure The human PRL (hPRL) and rat PRL (rPRL) genes are approximately 10 kb long from the 5¢ transcription initiation site to the poly A addition site at the 3¢ end and both consist of 5 exons separated by 4 large introns [60,61] (Fig. 4.2). Exon 1 consists of 56 bp that code for a portion of the 5¢ untranslated portion of the messenger RNA (mRNA) and the initial 28 aminoacids of the signal peptide. Exon 2 comprises 56 base pairs coding for the rest of the signal peptide and 120 base pairs coding for the first 40 amino acids of PRL itself. Exons 3 and 4 code for the amino acid sequences 41–76 and 77–136 respectively. Finally, Exon 5 codes for amino acids 137–199, the termination triplet TAA

FIGURE 4.2. details, see text.

and 118 base pairs before the AATAAA polyA signal [61]. The introns have sizes of 1.3, 1.5, 1.9 and 2.0 kb for A through D respectively [44]. The TATA sequence (Goldberg–Hogness box) is located 29 bp upstream of the start site of transcription [61]. The mature PRL mRNA that results after nuclear processing of the original heteronuclear RNA is only about 1 kb long and codes for a 227 amino acid sequence which includes an initial 28 amino acid signal peptide as well as the 199 amino acid structural peptide [19,61,62]. Almost 50% of the protein has an a-helix structure and the mature protein folds in a four-helix bundle similar to GH [63]. The 5¢ flanking region of the PRL genes contains an initial 500 base pairs region with considerable sequence homology between human, rat and bovine species, indicating that possible important regulatory elements are conserved there [61,64]. In the 5¢ flanking region, five highly conserved (79–90% homology between hPRL and rPRL) sequences for PRL are not found in hGH or rGH, indicating several specific sites that might be specific for PRL gene regulation in this regard [61].

5¢ Flanking Region and Tissue Specific Enhancers Detailed analyses of the 5¢ flanking region have been confined to the rat, using such models as the GH3, GH1, and GH4C1 tumor cell lines (Fig. 4.3). Preliminary analyses of the bovine PRL gene suggest common features in the 5¢ flanking regions across species [65], so that the findings to be discussed are likely also present in the human gene, with modifications.

Processing of the human prolactin gene. The exons are denoted by the numbers (1–5) and the introns by letters (A–D). For

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FIGURE 4.3. Detailed analysis of the 5¢ flanking region of the rat prolactin gene. ERE, estrogen response element; TRE, thyroid response element. 1P, 2P, 3P, and 4P refer to proximal binding sites for Pit-1; 1D, 2D, 3D and 4D refer to distal binding sites for Pit-1. For details see text.

There are two regions responsible for tissue-specific transcription activation, i.e. that allow expression of the PRL gene solely in the pituitary lactotrophs and not in other tissues; a distal enhancer element in the -1720 to -1580 bp region accounts for 98% to 99% of this activity while 1% to 2% of this activity is located at a more proximal promoter element at -422 to +33 [66,67]. Within these elements there are A,T—rich consensus sequences with a core of T/A T/A T/A ATANCAT that is critical and serves as a binding site for a tissue-specific transcription activator protein, termed Pit-1 [66,67]. Within the proximal region there are four binding sites for Pit-1, referred to as 1P (-38 to -62), 2P (-115 to -130), 3P (-142 to -162), and 4P (-180 to -200); similarly within the distal region the four binding sites for Pit-1 are referred to as 1D (-1550 to -1578), 2D (-1619 to -1636), 3D (-1645 to -1670), and 4D (-1694 to -1718) [67]. Although the distal and proximal regions are each capable of directing cell-specific expression of the PRL gene, they also appear to act synergistically [67]. The transcription enhancement of the PRL gene by the Pit-1 activator protein can be modified by other factors, such as TRH, epidermal growth factor (EGF), cAMP [68,69], and estrogens [54]. Others have also reported similar findings with respect to these distal and proximal sites [70]. Maurer et al. [71] have also found the tissue specific factor to bind to the sequence -1666 to -1659. Lemaigre et al. [72] demonstrated three pituitary specific protein binding sites in the proximal 217 bp of the PRL promoter in the 5¢ flanking region of the human PRL gene. Pit-1 consists of 291 amino acids with a molecular mass of 32.9 kDa [67]. Human Pit-1 cDNA has 85% DNA homology and greater than 95% amino acid sequence homology with the bovine, rat and mouse sequences

[46,73]. Pit-1 is part of the POU-domain gene family in which the DNA binding domain consists of a POUspecific domain and a POU-homeodomain [74]. Pit-1 is expressed in lactotrophs, somatotrophs and thyrotrophs and is the critical cell-specific transcription factor for activating expression of the PRL, GH, and TSH-b genes and for their respective cell development and proliferation [75,76] (see above).

Estrogen Also in the distal enhancer region lies the estrogen-response element, which has been localized to the position -1582 to -1569 [75,77,78]. This region binds the estrogen receptor and is required for the enhanced transcription of the PRL gene due to estrogen stimulation. The base sequence in this region of the rPRL gene represents an imperfect palindrome similar to the perfect conserved palindrome, GGTCANNNTGACC, which has been found to be necessary for the estrogen responsiveness of the vitellogenin gene in other tissues [75,78]. A second possible estrogen response element may also exist at -1722 to -1709 [77,79]. As noted above, estrogen regulates PRL gene transcription only if the PRL promoter is bound by Pit-1 [54].

Thyroid Hormone Two areas of the 5¢ flanking region have been found to be involved in the regulation of PRL gene transcription by thyroid hormones. The region -1565 to -1530 appears to be a stimulatory thyroid hormone response element that is distinct from the estrogen response element and can bind to the thyroid hormone receptor [80]. Day and Maurer [80] demonstrated by mutation analysis that the specific sequence

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of GGTCA at position -1555 to -1551 was shown to be critical and is the only portion of this region which displays sequence identity to a portion of the consensus sequence that has been derived for thyroid hormone response sequences [81]. A proximal region, -176 to -38, is involved in the ability of thyroid hormone to inhibit PRL gene transcription, but the thyroid hormone receptor does not bind to this region and it is likely that this inhibitory activity is indirect via some other trans-acting protein [80,82].

Intracellular Calcium Changes in cytosolic free calcium concentrations mediate the lactotroph responses to a number of extracellular stimuli (see below). A calcium response element has been demonstrated to reside within the initial 174 base pairs of the 5¢ flanking sequence [83]. The effects of intracellular calcium on enhancing PRL gene transcription may, in part, be mediated by the phosphorylation of a nuclear matrix protein kinase, topoisomerase II, by a calcium-calmodulin dependent protein kinase [84]. Four topoisomerase II recognition sequences were found in the 5¢ flanking region, at regions +1500 to +1600, -42 to -63, -1200 to -1300 and -2900 to -3000 base pairs with respect to the transcription start site [84]. Site 2, at -42 to -63 base pairs upstream from the transcription start site is within the region cited above as containing a calcium responsive element. Day and Maurer [85] have also found that both the proximal and distal enhancer regions are involved in mediating the transcriptional response to calcium. As discussed below, the activation of the phosphotidyl inositol pathway is probably a major route of stimulation of PRL gene transcription via an increase in intracellular calcium and the phosphorylation of protein kinase C. Direct stimulation of protein kinase C via phorbol ester causes PRL gene transcription via a regulatory region located at -74 to -30 of the 5¢ flanking region [68].

Glucocorticosteroids The region between -200 and +75 is necessary for glucocorticoid down-regulation of PRL gene transcription in the rat [77]. Sakai et al. [86] have provided evidence in the cow that the glucocorticoid response element (GRE) is a compound element with a constitutive enhancer-like component that activates transcription when glucocorticoids are not present and a sequence in which receptor binding reverses this enhancement. This negative GRE appears to lie between -310 and -152 of the 5¢ flanking region of the bovine PRL gene.

Cyclic AMP The sequence mediating the stimulatory effect of cAMP lies between -127 and +73 [70]. The sequence CCCCTCCC

lying between -78 and -71 in the rat gene has been shown to be a cAMP responsive element which binds to a transactivating factor termed Activator Protein 2 (AP-2) [87–89]. AP-2 has a molecular mass of 50–52 kDa [88]. In addition to this AP-2 binding region, the -97 to -84 region contains the sequence TGACGAAA which is similar to a cAMP response element (CRE) and may bind a CRE binding protein (CREB) [89]. There is also evidence that cAMP induced stimulation of PRL gene transcription may actually occur via the release of an inhibitory element downstream of -127 [70] or via intracellular calcium-dependent events [90]. Stimulation of PRL transcription by cAMP may also occur by augmenting the tissue specific enhancer element, Pit-1, discussed above [88,90].

PROLACTIN GENE REGULATION PRL secretion is influenced by a number of factors. Some of these factors, such as estrogen, have a direct action on the gene to alter transcription. Others, such as TRH and DA, act by binding to specific receptors on the lactotroph membrane, thereby activating intracellular second messenger signal transduction systems. This section will review these direct acting substances and these intracellular signal transduction systems (Fig. 4.4).

Estrogen The hormonal milieu of pregnancy has long been known to be stimulatory to lactotroph growth, causing hyperplasia of the normal pituitary lactotrophs [23,24] and enlargement of prolactinomas [91]. In early animal studies, it was shown that the PRL surge that occurred during proestrus was triggered by the prior estrogen peak [92] and that estrogen administration caused an increase in PRL synthesis [93–95]. Estrogen administration in vivo or when added to cell cultures results in an increase in lactotroph mRNA levels, paralleling the increase in PRL biosynthesis [96–101]. This stimulation of PRL gene transcription is biphasic. The first phase begins within 30 minutes of exposure and peaks within 1 hour, and involves the direct binding of estradiol to the nuclear estrogen receptor and subsequent binding of the estrogen-receptor complex to the specific estrogen response element discussed above. The second phase begins within 6 hours, continues for 24 to 48 hours and is indirect, in that it involves synthesis of a protein intermediate, as shown by this effect being blocked with protein synthesis inhibitors such as cycloheximide [100,102,103]. This second phase may be due to a modulation of the inhibitory effect of DA on PRL gene transcription, which itself is mediated by a decrease in intracellular cAMP [103,104]. Estrogen administration also stimulates mitotic activity, DNA synthesis and lactotroph proliferation [105–108].

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FIGURE 4.4. Signal transduction mechanisms involved in prolactin regulation. ER, estrogen receptor; TR, thyroid hormone receptor; GR, glucocorticoid receptor. In cytoplasm: ER, endoplasmic reticulum. For details, see text.

Thyroid Hormone

Adenyl Cyclase and Cyclic AMP

Thyroid hormones have long been known to suppress PRL synthesis and secretion from pituitary cell cultures [109]. Maurer [110] demonstrated a concomitant decrease in PRL mRNA levels, indicating an effect on PRL gene transcription. The roles of direct binding of the thyroid hormonereceptor complex to the distal thyroid hormone response element vs indirect effects via trans-acting factors at the more proximal site mentioned above in this inhibitory effect of thyroid hormone still need to be established.

Increases in intracellular cAMP result in an increase in PRL gene transcription and hormone synthesis and release in some cell systems but not others [103,114–116]. Agents which directly activate adenyl cyclase, such as forskolin, stimulate PRL gene transcription [116,117]. As discussed above, there may be both proximal and distal regions of the 5¢ flanking region that are responsible for the response to cAMP. It is likely that the effects of cAMP on gene transcription occur via the activation of a cAMP-dependent protein kinase and resultant phosphorylation of a specific DNA binding protein (AP-2, see above) associated with the cAMP response element (AP-2 binding site) of the PRL gene [118] or perhaps also via modulation of the effects of Pit-1 [70]. There are likely additional enzymes and proteins similarly phosphorylated that may be involved in eventual secretion of the mature PRL molecule. Furthermore,

Glucocorticoids Glucocorticoids exert an inhibitory action on PRL synthesis [111–113] and gene transcription. As discussed above, the inhibitory action may actually be a turning off of a constitutive enhancer-like component [86].

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protein phosphatases which interrupt signal transduction by decreasing phosphorylation are also important [119]. DA, the primary PRL inhibitory factor (PIF), acts through the D2 DA receptor, in part by inhibiting adenyl cyclase and decreasing intracellular cAMP [120,121]. Maurer found that adding monobutyryl cAMP to ergocryptine-pretreated cells resulted in a stimulation of PRL gene transcription [114] and Anderson et al. [122] were able to reverse the inhibitory effect of DA by adding forskolin, which increases intracellular cAMP levels. However, other experiments have shown that maintenance of intracellular cAMP levels does not abolish the inhibitory effects of DA on PRL release [123,124], so that other pathways may also mediate some of the inhibitory effects of DA. The intracellular phosphoinositide/calcium pathways may well be more important in mediating the inhibitory effect of DA on PRL release (see below). Vasoactive intestinal peptide (VIP) stimulates PRL release primarily through stimulation of adenyl cyclase and the production of intracellular cAMP [125–127]. However, it is unlikely that other secretogogues work through this pathway [128].

Phosphoinositide/Arachidonate Pathways A number of factors affecting PRL secretion act through the phosphoinositide pathways, including TRH, angiotensin II and DA [128]. As in other systems, after hormonereceptor interaction, coupled via a G protein, the initial step is the phospholipase C hydrolysis of phophatidylinositol 4,5biphosphate to yield inositol 1,4,5-triphosphate (IP3) and diacylglycerol (DAG). Stimulation causes an increase in IP3 which then causes a rapid release of calcium from the endoplasmic reticulum and an activation of membrane calcium channels to increase transport of extracellular calcium, resulting in a net increase in intracellular calcium content. Changes in intracellular calcium result in a number of further changes (see below), including, with DAG, activation of protein kinase C, which then acts by phosphorylating other proteins. The role of the phosphoinositide-protein kinase C pathway in the regulation of PRL secretion has been investigated in great detail. Protein kinase C is located in the normal pituitary [129], various PRL-secreting rat pituitary lines [130,131], and human pituitary tumors [132]. Activation of protein kinase C by phorbol ester results in PRL gene transcription with an increase in PRL mRNA levels and eventual PRL release from pituitary cells in vitro [133–137]. The mechanism mediating this PRL gene transcription is probably the protein kinase C phosphorylation of intracellular proteins [138,139] that, in turn, act as transacting factors on DNA regulatory elements in the proximal promoter region of the 5¢-flanking region of the PRL gene [140]. The phosphoinositide-protein kinase C pathway is a primary mediator of the stimulatory effects of TRH on PRL

gene transcription and release. TRH results in a rapid, phospholipase C-mediated hydrolysis of phosphotidylinositol with generation of DAG and IP3 [141,142]. The released DAG stimulates protein kinase C, resulting in phosphorylation of intracellular [139] and specifically nuclear, chromatin-binding proteins [138]. DAG probably mediates the second phase of the PRL response to TRH that occurs between 5 and 60 minutes after stimulation [143]. The increase in IP3 stimulates release of intracellular calcium stores [136,144], another mechanism involved in PRL gene transcription and release (see below). In various in vitro preparations, angiotensin II, neurotensin and bombesin have been found to stimulate PRL release in a similar fashion [128]. Studies of the interaction of DA with the pituitary D2 DA receptor and the phosphoinositide-protein kinase C pathway have yielded conflicting results, in part due to differences in cell preparations and experimental designs (reviewed by Lamberts and MacLeod [128]). Because chronic D2 receptor activation results in an attenuation of the phosphorylation of phosphotidylinositides [145], Lamberts and MacLeod [128] have concluded that DA’s inhibitory activity on PRL secretion occurs via this effect on phosphorylation rather than through an effect on phospholipase C. A second pathway involving membrane phospholipids is the arachidonate pathway, although the specifics of this pathway have not been fully elucidated. Stimulation of membrane-bound phospholipase A2 by TRH, angiotensin II, neurotensin and other direct stimulators results in PRL and arachidonic acid release from pituitary cells in vitro [146,147]. This effect of arachidonic acid release is blocked by inhibitors of phospholipase A2 [146] and by DA [148]. Arachidonate itself can liberate PRL in such cultures, an effect blocked by inhibitors of the conversion of arachidonate to its metabolites [149]. As only inhibitors of the lipoxygenase and epoxygenase pathways of arachidonate metabolism have an inhibitory effect of PRL release, it is likely that one or more metabolites in these pathways are the proximal effectors of PRL release [150]. The fact that arachidonate metabolism inhibitors have a much greater effect on inhibiting PRL release in basal conditions compared to the stimulated state suggests that these eicosanoids may be more likely to mediate paracrine or autocrine modulations of secretory mechanisms, rather than to function as intracellular messengers [150]. The effect of arachidonic acid in increasing PRL release is primarily via an increase in calcium influx rather than by mobilizing intracellular calcium stores [151].

Intracellular Calcium As discussed above, activation of the phosphoinositide pathway results in a mobilization of intracellular calcium stores and an influx of extracellular calcium. The resultant increase in intracellular free calcium results in PRL gene

Chapter 4

transcription, PRL synthesis and release. The mobilization of intracellular calcium is rapid but transient, resulting in a high amplitude burst of PRL release [128,152]. The stimulation of calcium uptake via calcium channels is more prolonged and proceeds at a lower rate, resulting in sustained PRL release [128,152]. As mentioned above, calcium interacts directly with DAG to activate protein kinase C. Calcium also binds to topoisomerase II which then interacts with specific DNA binding sites and to calmodulin, which binds to a number of different intracellular enzymes [84]. Preston et al. [153] demonstrated that the increase in intracellular PRL mRNA is due not only to an increase in gene transcription but also to a posttranscriptional mechanism involving the posttranslational modification of a stable protein which then regulates PRL mRNA stability. Blockade of calmodulin activity with penfluridol or selective antagonists such as W7 results in a decrease in PRL release [154,155], whereas the calcium ionophore, A23187, stimulates PRL release [155]. Furthermore, A23187 stimulates intracellular cAMP accumulation and W7 blocks both the A23187 stimulation of intracellular cAMP accumulation and PRL release [155]. Thus mobilization of intracellular calcium may have effects on both protein kinase C and cAMP. Calcium channel antagonists, such as verapamil, diltiazem, nifedipine, nimodipine, pimozide, penfluridol and the negative enantiomer of Bay K8644 inhibit, and agonists, such as the positive enantiomer of BAY K8644, stimulate PRL gene transcription, synthesis and release in in vitro preparations [156–162]. The effects of manipulation of intracellular calcium levels on the stimulatory and/or inhibitory effects of a number of physiologic PRL regulators have been assessed. Agents which mobilize intracellular calcium stores, such as A23187 and ionomycin, do not add to the PRL releasing effects of TRH, whereas BAY K8644, which stimulates calcium channels, does show an additive effect [159,163], suggesting that the effect of TRH on the first phase of PRL release is primarily through release of calcium stores rather than by stimulating calcium channels [162–166]. However, when these experiments were examined with respect to the action of TRH on PRL gene transcription and PRL synthesis, none of these agents were additive with TRH. Thus,TRH appears to cause rapid release of PRL via activation of protein kinase C and calcium store mobilization but not calcium channel transport, and its effects on PRL gene transcription and PRL synthesis involve all three processes [159]. La Verriere et al. [159] postulate that the initial steps involve activation of protein kinase C and mobilization of intracellular calcium, which in turn control the activity of calcium channels. DA inhibits the PRL release induced by activation of calcium channels by BAY K8644 [157,160] and maitotoxin [123] and by mobilization of intracellular calcium stores by A23187 [167]. Conversely, the calcium channel blockers nimodipine, verapamil and diltiazem antagonize the inhibitory effect of DA in vitro [168]. DA administration blocks

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the rise in intracellular calcium concentrations induced by TRH and angiotensin II [169], and DA withdrawal from the medium causes a rapid influx of calcium [170]. These findings are consistent with an inhibitory action of DA on calcium influx via calcium channels. Maitotoxin, which activates calcium channels, also blocks the DA induced inhibition of PRL secretion from lactotrophs in vitro [123]. Thus, at least part of the inhibitory action of DA on PRL secretion appears to be mediated by effects on intracellular calcium concentrations. In humans, administration of the calcium channel blocker verapamil causes an increase rather than a decrease in serum PRL levels [171,172] and does not inhibit the TRHinduced PRL rise [172]. This was shown to be due to a decrease in tuberoinfundibular generation of DA [173]. Because dihydropyridine and benzothiazepine calcium channel blockers had no effect on PRL levels in humans, it was hypothesized that the effect of verapamil is via action at N-type calcium channels present in neuronal tissues [173].

HORMONE BIOSYNTHESIS The primary translation product of the human PRL gene is 227 amino acids in length, consisting of a 28 amino acid signal peptide and a 199 amino acid hormone with a molecular mass of 23-kDa [19]. The structure predicted by the mRNA sequence delineated by Cook et al. [19] differs at eight positions from the early sequence of Shome and Parlow [18], which was determined by standard sequencing techniques. There are three disulfide bonds between cysteines at positions 4 and 11, 58 and 174 and 191 and 199, giving rise to small loops at the amino and carboxy terminal ends and a large loop in the middle (See Fig. 4.2). There is considerable heterogeneity in the final PRL product, depending upon variations in the degree of posttranslational modifications, which include cleavage, polymerization, glycosylation, phosphorylation and degradation (Table 4.1).

Cleavage Products Small amounts of 16 K and 8 K cleavage products are present in the normal pituitary and plasma of humans [174] as well as rats [175] and mice [176]. In the rat, the 16 K fragment has greater mitogenic activity than the intact hormone [177], but this has not yet been determined for hPRL. Several PRL-like proteins that are only a few amino acids shorter than the 23 K form and have a molecular weight of about 21 K are present in the rat pituitary. These smaller variants arise from carboxy (COOH)-terminus proteolytic clipping during storage and not from transcriptional modifications resulting from alternative splicing of the PRL pre-mRNA, as only a single 23 K band is found with in vitro translation systems [178–181]; these have not yet been described in the human.

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Table 4.1. of prolactin

Hypothalamic–Pituitary Function

Posttranslational processing and other variants

Cleavage 23K 21K (?) 16K 8K

Polymerization Monomer—80–90% Dimer—8–20% Polymer—1–5%

Glycosylation Circulating—50–100% Pituitary—13–25%

Phosphorylation

Decidual prolactin Lymphoblastoid prolactin

different types of glycosylation products have been described [197]. The glycosylated variants account for 13% to 25% of pituitary PRL and 50% to 100% of circulating PRL [196–199]. Glycosylated PRLs are less immunoreactive in the routine PRL RIA [196], bind less well to the PRL receptor [200] and are less bioactive, using a variety of bioassays [197,201,202]. On the other hand, Young et al. [203] found glycosylated porcine PRL to have greater lactogenic and binding affinity, despite decreased mitogenicity compared to nonglycosylated PRL. Interestingly, during pregnancy, subsequent lactation, and pathologic hyperprolactinemic states, when there is a high PRL secretion rate, the relative amount of nonglycosylated PRL increases compared to other times [199,201].

Placental variants

Phosphorylated Forms Larger Molecular Weight Forms About 80% to 90% of PRL extractable from pituitaries and in serum is monomeric, 8% to 20% is dimeric (molecular mass 45–50 kDa) and 1% to 5% is polymeric [182–186]. The dimeric and polymeric forms are linked by disulfide bonds [184] (see below) and can be largely dissociated under reducing conditions or with digestion with chymotrypsin [184,185]. Polymerization occurs primarily within storage granules and in stimulated lactotrophs PRL is synthesized and discharged rapidly, bypassing polymerization [187]. However, with stimulation the stored polymeric forms are released into plasma [188]. These larger molecular weight polymers have decreased binding to receptors and display decreased bioactivity in a variety of receptor assays [183,184] but have normal bioactivity in the Nb2 lymphoma cell bioassay (see below) [186]. Using highly purified, monomeric human PRL infused into normal male subjects, Molitch et al. [189] showed that some conversion of monomer to dimer also occurs in plasma. A number of patients have been described with elevated basal serum PRL levels but normal reproductive function who were found to have markedly elevated proportions of polymeric PRL, presumably thereby resulting in less PRL bioactivity [190–194]. However, in many such reported cases the proportion of monomeric PRL in the blood is still elevated. Preliminary screening of sera suspected of having a high proportion of polymers can be done by precipitating the large complexes with polyethylene glycol, but gel filtration chromatography remains the best procedure for quantitating the amount of polymers [195]. In clinical practice, screening for polymers is rarely carried out.

Glycosylated Products Glycosylation of human PRL occurs [196]. The carbohydrate unit is linked to the asparagine at position 31 and the complex has a molecular weight of about 25 K [196]. Two

PRL can be phosphorylated within secretory granules in rat and bovine lactotrophs [181,204,205]. These secretory granules appear to contain a PRL kinase, although the specifics of this remain to be determined. The role of this phosphorylation in secretion, whether phosphorylated forms circulate, and whether this process occurs in humans are unknown.

Placental Variants In mice and rats, there are a variety of PRL-like proteins produced by the placenta in addition to placental lactogen [206,207]. These PRL-like proteins are secreted at different times during gestation by the placenta and thus may have different functions, including alterations in blood vessel formation, hematopoiesis, and lymphocyte function [206,207]. No such proteins have been identified in the human and human placental lactogen is much closer in structure to GH than it is to PRL.

Decidual Prolactin PRL levels in maternal blood rise throughout gestation and are of pituitary origin [208,209]. However, PRL concentrations in amniotic fluid are 10–100 fold higher than either maternal or fetal blood levels [208]. Early biochemical studies found that amniotic fluid PRL was identical to serum PRL biologically, chemically and immunologically [210,211]. Friesen et al. [212] first showed that cultured human chorion released PRL and subsequently Riddick and Kusmik [213] found that cultured human decidual cells released it. Finally Golander et al. [214] documented that cultured human chorion-decidual explants synthesize and release a PRL that was shown to be identical to pituitary PRL. It was further documented that the decidua produced PRL [215], and that its bioactivity was equal to pituitary PRL [216]. Additional studies have shown that decidual PRL mRNA hybridizes with a pituitary PRL

Chapter 4

cDNA probe and Northern blot analysis shows that the 2 PRL mRNAs are indistinguishable, except for four silent nucleotide differences and the decidual PRL gene being about 150 nucleotides longer in the 5¢ untranslated region [217–219]. Similar to pituitary PRL, decidual PRL is also glycosylated [220] and some of the glycosylated PRL is bound to immunoglobulin by disulfide bonds in the amniotic fluid [221]. Similar rat decidual PRL-like substances have been reported that are structurally different from rat pituitary PRL but bind to PRL receptors and stimulate steroidogenesis by the corpus luteum [222–224]. The regulation of decidual PRL production differs from that of pituitary PRL (detailed below). DA agonists [225] and antagonists [226] given to the mother respectively decrease and increase maternal serum PRL levels but have no effect on amniotic fluid PRL levels. Similarly, DA, bromocriptine, and TRH have no effects on the decidual production of PRL in vitro [214]. Decidual PRL production is increased by progesterone and progesterone plus estrogen but not estrogen alone [218,227]. In 1988, Golander et al. [228] reported that a purified, 23.5-kd placental protein could stimulate the synthesis and release of PRL from decidual cultures. Insulin, through the insulin receptor, insulin-like growth factor I (IGF-I), through the IGF-I receptor and relaxin, a third peptide related to insulin and IGF-I have all been reported to stimulate synthesis and release of PRL [218,229,230]. The function of decidual PRL remains obscure, although there is some evidence that it may contribute to the osmoregulation of the amniotic fluid [231], fetal lung maturation [232] and uterine contractility [233].

Lymphoblastoid Cell Prolactin A human clonal lymphoblast cell line, IM-9-P3, has been established which produces a PRL that is identical to pituitary PRL in all aspects [234], including the sequence of their mRNA’s [235]. There are differences from pituitary but not decidual PRL, however, in the 5¢ untranslated region of the gene, suggesting that regulatory mechanisms may be different [235]. Indeed, preliminary studies have shown that PRL production by this cell line can be decreased by dexamethasone but other substances known to affect pituitary PRL secretion, such as estradiol, thyroxine, TRH and VIP, have no effect [236]. The similarities in the structure and regulation of the IM-9 and decidual PRL genes suggest that studies of the IM-9 cell line may prove fruitful in discerning the mechanisms regulating decidual PRL production, but it is doubtful that such studies will contribute much to our understanding of pituitary PRL regulation. HORMONE SECRETION—BIOCHEMISTRY Following synthesis on the rough endoplasmic reticulum, PRL is packaged into secretory granules in the Golgi and

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PRL release occurs via exocytosis [237,238]. Nicoll and Swearingen in the early 1970s [239,240] demonstrated the existence of two pools of PRL within the rat lactotroph cell, one turning over rapidly and the other turning over slowly. Newly synthesized PRL is preferentially released compared to older, stored PRL in response to some stimuli and constitutes the rapidly turning over pool [238,241–243]. However, other stimuli, such as TRH, result in a preferential release of older, stored PRL [238]. Walker and Farquhar [238] found that these two types of secretion, rapid and slow, occurred not so much due to differences in the type of stimulation for a given cell but due to functional heterogeneity of the cells, so that some cells synthesize and secrete PRL rapidly while others secrete more slowly (see also the discussion above about the hemolytic plaque assay). Much of the storage pool of PRL in the pituitary appears to exist in a high molecular weight, disulfide-bonded, poorly immunoreactive polymeric form that is converted to a releasable, immunoreactive monomeric form within the secretory granule when processed for release [244]. This conversion involves a thio-disulfide interchange mechanism and can be stimulated by reduced glutathione and decreased by aminothiols such as cysteamine [244,245]. Treatment with cysteamine, in fact, reduces the amount of PRL detectable within cells using immunoprecipitation and RIA, but not when electrophoresis alone is used for detection [246,247]. These changes appear to explain a previous finding in rats that stimulation resulted in a “depletion” of the storage form of PRL in the pituitary with its “transformation” into the readily releasable, immunoreactive PRL [248]. Cysteamine has also been shown to reduce PRL release from pituitaries in humans and its therapeutic use in patients with hyperprolactinemia has been suggested [249], although its action on disulfide bonds in other peptides, such as somatostatin, would appear to preclude such use [250].

Measurement of PRL RIA

RIAs for PRL were initially developed for the rat [6], sheep [7] and cow [8]. After studies finally showed that in humans PRL and GH were separate molecules (see above), RIAs were developed for human PRL [16,17]. Most RIAs are double antibody assays, using as standard PRL that is either of commercial origin or a preparation available from the National Hormone and Pituitary Program in the U.S.A. or the National Institute for Biological Standards and Control in the UK. Because of the above noted differences in the kinds of PRL that exist in the pituitary and serum, it is not surprising that antibodies raised against one preparation of PRL may bind PRL differently from antibodies raised against a different preparation of PRL. Furthermore, PRL preparations for iodination and standards differ from one laboratory to another and therefore some means of standardizing one assay with another is

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important. Although antibodies still differ, at least the standards used can be standardized against a reference preparation, and this is usually done. First, second and third International Reference Preparations (IRPs) have been established by the National Institute for Biological Standards and Control in London [251,252] and similar reference preparations are made available by the National Hormone and Pituitary Program in Baltimore. Unfortunately, the supply of suitable standards does not preclude considerable variation from RIA to RIA in different laboratories in the measurement of PRL levels in both research studies and clinical evaluations of patients. In a survey of 10 different laboratories, the mean intraassay coefficient of variation was 9.6% but ranged from 5.6% to 25.3% and the mean interassay coefficient of variation was 7.9% [253]. Over the past decade, two-site immunoradiometric assays (IRMA) and chemiluminometric assays (ICMA) have come into wide use because of improved sensitivity and precision and short incubation periods. However, in patients with large prolactinomas with very high PRL levels, the very high level of PRL may saturate the antibodies, preventing the formation of the PRL-antibody sandwich, leading to loss of labeled antibody and a falsely low value [254,255]. St. Jean et al. [254] noted this high end “hook effect” in 5.6% of 69 patients thought to have clinically nonfunctioning adenomas. Therefore, in patients with large macroadenomas, PRL assessments should always be performed in both undiluted and 1 : 100 diluted serum to exclude the “hook effect” when two-site assays are used. Bioassays

The pigeon crop sac assay for PRL was the first assay used and was the standard until the late 1960s [1,3,4]. Although the procedure was refined and quantified substantially by Nicoll [4], it still proved cumbersome. Kleinberg and Frantz subsequently developed a highly sensitive bioassay which involved grading histologic changes in mouse mammary tissue that had been cultured with varying concentrations of PRL or serum samples [256]. It was with this assay that Kleinberg and Frantz first demonstrated that PRL is a separate hormone from GH in human blood [15]. This assay was further refined by Turkington [257], with quantitation of casein production by the mammary tissue being the endpoint measured. A radioreceptor assay using rabbit mammary gland membranes has been developed [258] which, as with other radioreceptor assays, essentially measures receptor binding but not true bioactivity. The most commonly used bioassay now is the Nb2 node rat lymphoma cell assay. In this assay cultured lymphoma cells replicate in response to added GH and/or PRL but not other hormones, with a sensitivity down to 0.1 ng/ml [259]. Using this assay, Tanaka et al. [260] and Rowe et al. [261] found bioassay/RIA ratios with a range of 0.53– 1.56 in sera from normal and hyperprolactinemic patients. As discussed above, in some patients with high amounts of polymeric PRL circulating in their blood, this ratio is

normal, whereas it is decreased in other bioassays. Thus this bioassay measures only one aspect of PRL function and when detailed assessments of biological function are necessary, more than one bioassay may be required. Frawley et al. [262] have revived the mammary gland casein production assay, achieving much better sensitivity. In this assay, they use a monolayer dispersed cell culture of rat mammary glands and then detect PRL induced casein production by a variant of their hemolytic plaque assay using casein antibodies, protein-A coated ovine RBC’s and complement, achieving a sensitivity of <1 pg/ml [262].

Physiology Metabolic Clearance and Production Rates of Prolactin

The metabolic clearance rate (MCR) and production rate (PR) of PRL have been determined by two methods: the infusion to equilibrium method using radioactively labeled PRL, and the infusion of exogenous hPRL into individuals in whom endogenous PRL production had been suppressed with a DA infusion. In two studies using the former method, the MCRs were found to be 46 ± 4 and 40 ± 6 mL/min/m2 [263,264] and with the latter method it was found to be 71 ± 19 mL/min/m2 [189]. The calculated PRs using the labeled PRL method were 200 ± 63 and 536 ± 218 mg/day/m2 and 802 ± 377 mg/day/m2 using the hPRL infusion method. The calculated disappearance half-lives varied from 26 to 47 minutes in these three studies. In another study the disappearance half-life was estimated to be 43.6 ± 11.2 minutes as determined from the rate of fall of PRL with individual secretory episodes [265]. The fate of PRL is unclear. Studies in patients with chronic renal failure have shown the MCR to be reduced by 33% [264] but Bratusch-Marrain et al. [266] found no hepatic uptake of PRL in man, in contrast to the liver’s being responsible for about 50% of total GH clearance in man and playing a substantial role in the uptake of PRL in rabbits [267]. Hormone Secretion Patterns

PRL is secreted episodically (Fig. 4.5). There is an innate pulsatility to pituitary PRL secretion with an interpulse interval of about 8 minutes, as determined by studies of plasma samples in rats in whom pituitaries were placed under the capsule of the kidney to escape hypothalamic regulation [268] and from media obtained from primate pituitaries cultured in vitro [269]. When plasma is sampled from normal individuals in whom hypothalamic function is superimposed upon this innate pulsatility, it becomes apparent that there are 4 to 14 secretory episodes per day. In two studies that used cluster analysis, 13 to 14 peaks per day in young subjects were found with a peak duration of 67 to 76 minutes, a mean peak amplitude of 3–4 ng/ml and an interpulse interval of 93 to 95 minutes [270,271]. Disinhibition caused by hypothalamic tumors causes an increase in

Chapter 4

FIGURE 4.5. Prolactin levels throughout the day in a single individual superimposed upon the range from five normal individuals. Note the episodic nature of secretion and the nocturnal rise.

basal PRL levels due to an increase in pulse amplitude and not pulse frequency [272]. There is an increase in the amplitude of the PRL secretory pulses that begins about 60 to 90 minutes after the onset of sleep; the secretory pulses increase with non-REM sleep and fall prior the next period of REM sleep [273]. Lowest PRL concentrations are found during REM sleep and highest concentrations are found during non-REM sleep [273]. When subjects are kept awake to reverse the sleep-waking cycle, PRL levels do not rise until sleep begins [274]. Thus, the diurnal variation of PRL secretion is not an inherent rhythm but depends on the occurrence of sleep. Interestingly, the diurnal variation of PRL with the sleep-induced rise persists despite other powerful physiologic influences such as breast-feeding [275]. Studies of PRL secretory patterns in twins show that there are both genetic and environmental influences on these patterns [276]. Weitzman et al. [277] noted small rises in PRL in the afternoon and evenings and they postulated that there may be a relationship to meals. Quigley and Ropert noted an increase in circulating PRL levels of 50% to 100% within 30 minutes of meals [278]. Subsequent work by Carlson related this increase to the amino acids generated from the protein component of the meals, phenylalanine, tyrosine and glutamic acid being the most potent in this regard [279–282]. Carlson et al. [282] have provided evidence that this stimulatory action of these amino acids is centrally mediated by showing that large neutral amino acids such as valine inhibit the transport of phenylalanine across the blood–brain barrier and blunt the stimulatory action of this amino acid. Changes in Prolactin with Age

PRL levels are elevated almost 10-fold in infants following delivery but then gradually decrease so that levels are normal by three months of age [283]. These high levels of PRL at birth are probably related to the stimulatory effect of high maternal estrogen levels. PRL levels are lowest between the ages of 3 months and 9 years and then rise modestly during

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FIGURE 4.6. Serum prolactin (PRL) concentrations measured serially at weekly intervals as a function of gestation (n = 4). The dashed line represents the linear regression and the solid line represents the second order regression. NP, nonpregnant PRL values. From Rigg et al. [209]

puberty to adult levels [283]. In some studies there is a gradual fall of basal PRL levels with age, but in other studies no changes with age have been found [271,283–290]. One recent study found that PRL levels in women gradually fall by about 50% over the first 18 months following menopause, but this fall is considerably less in women treated with estrogen replacement therapy [287]. Other studies have shown no change in PRL levels with hormone replacement therapy [288]. In hyperprolactinemic women, estrogen replacement therapy causes no change in PRL levels [289]. A recent detailed study showed that mean serum PRL concentrations were decreased by 55% in older compared to younger men, and that this decrease was due to decreased basal secretion as well as the amount of PRL secreted with each secretory burst but that there was no change in pulse duration or frequency or interpulse interval [290]. Changes in Prolactin Levels during the Menstrual Cycle

A number of investigations of the changes in blood PRL levels during the menstrual cycle have revealed that some, but not all, subjects have higher levels at mid-cycle and lower levels in the follicular compared to the luteal phase [208,253,285,291–294]. In most of these studies, no correlations were found between PRL and estradiol, progesterone, LH and FSH levels. However, some studies have shown that PRL and LH secretion are often synchronous in the luteal phase and that very small doses of GnRH can cause the secretion of both PRL and LH at this time [295]. Changes in Prolactin Levels during Pregnancy

Basal PRL levels gradually increase throughout the course of pregnancy (Fig. 4.6) [208,209]. This has generally been attributed to the stimulatory effect of the hormonal milieu of pregnancy, primarily estrogenic, on the pituitary

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lactotrophs. There is a gradual increase in the number of pituitary lactotrophs during pregnancy [23,24] and by term, PRL levels may be increased 10-fold to levels over 200 ng/ml [208,209,283]. These elevated PRL levels found at term prepare the breast for lactation. Changes in Prolactin Levels with Postpartum Lactation

Within the first 4–6 weeks postpartum basal PRL levels remain elevated in lactating women and each suckling episode triggers a rapid release of pituitary PRL resulting in a 3–5 fold increase in serum PRL levels, peaking about 10 minutes after the end of suckling [208,296]. This increase in PRL secretion is accompanied by a suckling-induced increase in pituitary PRL mRNA levels which provide for increased PRL synthesis [297]. Following termination of suckling PRL levels gradually fall to reach prenursing levels by about 3 hours after the beginning of the suckling episode [296,298,299]. Over the next 4–12 weeks, basal PRL levels gradually fall to normal and the PRL increase which occurs with each suckling episode decreases [296,300]. Eventually there is little or no rise in PRL with suckling despite continued milk production [208]. The actual PRL levels achieved during suckling do not correlate with the amount of milk produced [298]. The decreases in basal and stimulated PRL levels between 3 and 6 months postpartum are largely the result of decreased breast-feeding as formula is introduced into the baby’s diet. If intense nursing behavior is maintained, basal PRL levels remain elevated and postpartum amenorrhea persists (Fig. 4.7) [301–303]. Eighty minutes of nursing per day with a minimum of six nursing episodes will usually result in persistent hyperprolactinemia and amenorrhea [303]. However, the delay in the onset of menses is more associated with high suckling duration and frequency than with a particular level of PRL [303]. High intensity lactation-induced failure to ovulate and menstruate has been used as a method of contraception in a number of developing countries for many years [304–306]. Breast stimulation may cause an increase in PRL levels in some non-breast-feeding normal women but not in men [296] except for one study showing PRL increases in men after breast stimulation by their wives [307]. Changes in Prolactin Secretion with Stress

PRL has long been known to be one of the pituitary hormones released by stress, along with adrenocorticotropic hormone (ACTH) and GH. Neill initially demonstrated the release of PRL from rats with ether stress [308] while Noel et al. initially showed a similar response to physical stress in humans [309]. The stress-induced rise in PRL generally consists of a doubling or tripling of PRL levels and lasts less than 1 hour. This rise in PRL may be limited by the concomitant release of cortisol which suppresses further PRL release, as evidenced by the fact that stress applied to adrenalectomized rats results in a PRL increase that is sustained for as long as the stressful stimulus is applied [310,311]. In

FIGURE 4.7. Mean 24-hour prolactin levels as a function of number of minutes of nursing in the same 24-hour period. Each circle represents one woman. The same amount of nursing results in higher levels of prolactin in the first year than in the second year postpartum.
= <12 months postpartum;  = >12 months postpartum. r = 0.54, P < 0.02. From Stern et al. [303]

humans, prolonged critical illness does not cause a sustained elevation of PRL; rather there is a reduction in the pulsatile secretion with an overall lowering of levels [312]. The teleologic significance for these stress-induced changes in PRL is not clear. The neuroendocrine mediation of the acute stress response is probably multifactorial but does not include a decrease in dopamine [313]. Corenblum and Taylor [314] attempted to dissect out the neurotransmitter regulation of the PRL stress response in humans by administering various blocking agents immediately prior to surgery. Blockade of histamine H1 receptors using chlorpheniramine, serotonin receptors using cyproheptadine, and dopamine receptors using pimozide had little effect on the peak PRL level reached during surgery. Blockade of opiate receptors with high dose naloxone resulted in a significant blunting, but not complete inhibition of the PRL response. These studies imply that the endogenous opiate-like peptidergic pathways may play a role in the PRL stress response. On the other hand, in humans naloxone has generally not been found to be able to block the PRL response to hypoglycemia [315]. Abe et al. found that VIP antisera inhibited the etherinduced PRL rise in rats [316] and Kaji et al. [317] found that antisera to VIP and PHI were additive in this regard. Other factors including serotonin, oxytocin and possibly unidentified posterior pituitary PRL releasing substances may also be involved, but their relative roles remain obscure [313].

Chapter 4

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Hypoglycemia has been regarded as a form of stress, but whether it acts as a nonspecific stressor or has more specific effects is not clear. Among their tests of various types of stress, Noel et al. [309] showed that PRL did indeed rise with hypoglycemia. Woolf et al. [318] evaluated this effect systematically in normal men and women, finding that PRL levels increased by at least 10 ng/ml with a doubling of baseline levels in two-thirds of normal subjects, the maximal rise occurring between 40 and 90 minutes after the injection of intravenous insulin. 2-deoxy-d-glucose, which causes intracellular glucopenia, also causes a rise in PRL levels [319]. Studies using combinations of inhibitors of dopamine, histamine and serotonin suggest that at least some of the PRL response to hypoglycemia may be mediated by serotoninergic pathways [320]. Acute exercise has also been regarded as a form of stress and results in an acute, transient increase in PRL levels [309,321]. Although chronic, high-level exercise often results in menstrual disturbance, it is not associated with sustained hyperprolactinemia [322]. Effects of Thyroid Hormone Status on Prolactin Secretion

As noted above, thyroid hormones have a direct suppressing action on PRL gene transcription and hormone synthesis, so that in hypothyroidism this effect is decreased. As discussed below, in hypothyroidism TRH synthesis is increased, pituitary TRH binding is increased and pituitary VIP levels are increased. The result is that in hypothyroidism, PRL levels may be significantly increased. NEUROENDOCRINE REGULATION The hypothalamus exerts a predominantly inhibitory influence on PRL secretion through one or more PRL inhibitory factors (PIF) that reach the pituitary via the hypothalamic–pituitary portal vessels (Fig. 4.8). There are PRL-releasing factors (PRF) as well (Table 4.2). Disruption of the pituitary stalk leads to a moderate increase in PRL secretion as well as to decreased secretion of the other pituitary hormones.

PIFs Dopamine

In 1954, Everett [323] demonstrated that the luteotropic properties of the pituitary (due to PRL) were increased when pituitaries were transplanted to beneath the renal capsule, a site away from the regulation by the hypothalamus, thus demonstrating the predominance of the inhibitory component of hypothalamic regulation of PRL secretion. Pasteels [324] and Talwalker et al. [325] subsequently showed that hypothalamic extracts suppressed PRL secretion from pituitaries in vitro. At about the same time, Sawyer’s group [326,327] demonstrated an increase in PRL secretion in rats in vivo with use of catecholamine depleting agents such as reserpine or with DA receptor blockers, such as chlorpro-

FIGURE 4.8. Neuroendocrine regulation of prolactin secretion. GAP, gonadotropin-associated peptide; PHM, peptide histidine methionine; PIF, prolactin inhibitory factor; PRF, prolactin releasing factor; TRH, thyrotropin releasing hormone; VIP, vasoactive intestinal peptide.

mazine. In 1968 van Maanen and Smelik proposed that tuberoinfundibular DA (TIDA), released into the hypothalamic–pituitary portal vessels in the median eminence, was the physiologic PIF [328]. Shortly thereafter, the direct pituitary site of action for DA was confirmed by experiments in which it was demonstrated that DA could suppress PRL release from pituitaries in vitro [329–331] and Diefenbach et al. [332] demonstrated that the DA precursor L-dopa could suppress PRL secretion in intact and stalk-sectioned monkeys. A number of experiments have now firmly established that DA is the predominant, physiologic PIF. Gibbs and Neill [333] demonstrated that the concentration of DA found in the pituitary stalk plasma (about 6 ng/ml) was sufficient to decrease PRL levels in rats and was five to 10fold higher than levels found in peripheral plasma. Stimuli

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Table 4.2.

Substances affecting prolactin release

Stimulatory Thyrotropin-releasing hormone Vasoactive intestinal peptide Peptide histidine isoleucine Serotonin Opioid peptides Growth hormone-releasing hormone Gonadotropin-releasing hormone Oxytocin Vasopressin Histamine (H1) Bradykinin Angiotensin II Neurotensin Substance P

Inhibitory Cholecystokinin Bombesin Secretin Gastrin Galanin Calcitonin Calcitonin gene-related peptide Thymosin factor 5 Melatonin Other posterior pituitary factors (?) Platelet activating factor Epidermal growth factor a-melanocyte stimulating hormone Prolactin-releasing peptide

which result in an acute release of PRL usually also result in an acute decrease in portal vessel DA levels, including such stimuli as estrogen [334], cervical stimulation [335], and mammary nerve stimulation simulating suckling [336]. However, in many experiments it was found that the PRL increase obtained by simply reducing DA was considerably less than the elevation of PRL achieved by simultaneous stimulation by a PRF; similarly, the PRL level achieved with the simultaneous stimulation by a PRF with the reduction in DA is usually greater than that achieved by a PRF alone [337–339]. It is likely that in most physiologic circumstances that cause a PRL rise, such as lactation, there is a simultaneous fall in DA along with a rise in a PRF, such as VIP [316]. However, there may well be circumstances in which various PRF’s may stimulate PRL release with no concomitant lowering of DA levels, or DA may be lowered with no concomitant increase in a PRF. Newer work with mice in which the DA D2 receptor or DA transporter have been “knocked out” (KO) with inactivating mutations has confirmed these earlier studies that employed pharmacologic methods or lesioning. Thus, mice with the DA D2 receptor KO develop lactotroph hyperplasia without adenomatous transformation and sustained hyperprolactinemia [340,341]. DA action within the synapse terminates by DA reuptake by the DA-secreting neurons via the DA transporter. In contrast to the findings with the DA receptor KO, DA transporter KO mice have increased dopaminergic tone and lactotroph hypoplasia [342]. Although such mice have normal circulating levels of PRL, they cannot increase these levels with various stimuli and are unable to lactate [342]. Interestingly, the elevated DA levels in the DA transporter KO mice also depresses hypothalamic GH releasing hormone (GHRH) content resulting in decreased GH secretion and growth [342]. Although much of the direct work demonstrating DA in hypothalamic–pituitary portal vessels and the effects of DA

Dopamine Gonadotropin-associated peptide g-aminobutyric acid Somatostatin Acetylcholine

FIGURE 4.9. Time course of the serum prolactin response to the intravenous injection of 400 mg of synthetic thyrotropinreleasing hormone (TRH) in 36 normal men and 36 normal women. From Jacobs et al. [406]

on PRL release in vitro have been done in animals, it is clear that DA is the primary PIF in humans as well. Infusion of DA causes a rapid suppression of basal PRL levels [342–346] that can be reversed by metoclopramide, a DA receptor blocker [346]. Dopamine also blocks the PRL increments induced by such stimuli as hypoglycemia [347], arginine [348] and TRH [349]. Studies with low-dose DA infusions in humans have shown that DA blood concentrations similar to those found in rat and monkey hypothalamic–pituitary portal blood [333,350] are able to suppress PRL secretion [351–353]. Blockade of endogenous DA receptors by a variety of drugs, including phenothiazines, butyrophenones, metoclopramide and domperidone causes a rise in PRL

Chapter 4

[354,355]. Furthermore, blockade of the decarboxylation of l-dopa to DA by the dopa decarboxylase inhibitor carbidopa in the peripheral blood and median eminence causes an increase in PRL [356]. The axons responsible for the release of DA into the median eminence originate in perikarya in the dorsomedial portion of the arcuate nucleus and inferior portion of the ventromedial nucleus of the hypothalamus [357–359]. This pathway is known as the tuberoinfundibular DA (TIDA) pathway and DA axon terminals comprise about a third of all axon terminals in the zona externa of the median eminence [360]. Some TIDA neurons do not terminate in the median eminence but project to the posterior and intermediate lobes where their endings are found close to pituicytes, magnocellular axon terminals and precapillary spaces [361]. In some experiments, removal of the posterior lobe results in an increase in basal PRL levels that lasts for a few days [362]. However, the precise physiologic role of the posterior pituitary in inhibiting PRL secretion in the rat remains unknown [362]. There are no data in humans regarding such a role for the posterior pituitary in PRL regulation. The DA that traverses the TIDA pathway binds to the class of DA receptors referred to as D2 receptors [363–365] on the lactotroph cell membrane [366,367]. As discussed above, activation of this receptor results in: (i) an inhibition of adenyl cyclase with lowered intracellular cAMP levels; (ii) inhibition of phosphoinositide metabolism; and (iii) decreased intracellular calcium mobilization and inhibition of calcium transport through calcium channels. Some studies have also suggested that at least some of the inhibitory action of DA may be mediated by nitric oxide [368]. It has been proposed that these different actions of DA may actually be mediated by multiple similar D2 receptors that are produced by alternative RNA splicing [369]. The functional state of the D2 receptor is the high affinity form [364,367,370]. Although binding of DA to the D2 receptor appears to be sufficient to decrease PRL secretion mediated by the above pathways, DA can also be found intracellularly in association with PRL secretory granules [371–373]. The functional significance of this association of DA with the secretory granules is unknown. The inhibitory action of DA on PRL secretion is partially blocked by estrogen administration. This may be largely due to the direct action of estrogen on the estrogen response element of the PRL gene (see above). However, there may also be other mechanisms but with considerable interspecies differences. Estradiol is able to block the inhibitory action of dopamine on PRL release from rat lactotroph cells in vitro [374–376] but not from monkey lactotrophs [377]. In studies in humans, Judd et al. [344] found that the same dose of infused DA resulted in a greater suppression of PRL during the early follicular phase, when estrogen levels are low, compared to the late follicular or periovulatory phases, when estrogen levels are higher. PRL levels are also suppressed to a greater extent in women compared to men by the same dose of estrogen [343]. Estrogens

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result in a decrease in DA receptor numbers in rats [378] but the DA receptor population showed no sex related differences in a limited number of human pituitaries [367]. Although portal vessel DA levels are higher in female rats or in male rats treated with estrogen [379], the direct effects of estrogen at the lactotroph level result in a blunting of the inhibitory effect of DA and increased PRL production [104,105]. Gonadotropin Associated Peptide (GAP)

Whether DA alone can account for all of the PIF activity of the hypothalamus has long been a question. In the mid-1970s a number of groups found considerable PIF activity in rat hypothalamic extracts in which DA had been removed [380–383]. In 1985 Nikolics et al. [384] reported the PRL inhibiting ability of a 56 amino acid polypeptide that is in the carboxyterminal region of the precursor to gonadotropin releasing hormone (GnRH) and which they termed GAP. In pituitary cultures, GAP was found to inhibit PRL secretion at concentrations of 10-11 M, whereas DA was found to inhibit PRL secretion only at much higher concentrations (10-7 M) [384]. Subsequent experiments showed a 100-fold lower potency of GAP for PRL suppression but at still lower molar levels than for DA [385,386]. Furthermore, passive immunization of rabbits against fragments of GAP resulted in marked elevations of endogenous PRL levels [384]. In these experiments, GAP was also shown to have weak gonadotropin stimulatory properties, giving rise to the concept of inverse stimulation/inhibition of these hormones in various physiologic circumstances. Antibodies raised to a synthetic peptide constituting residues 40–53 of GAP colocalized with GnRH in secretory granules of nerve terminals in the median eminence of the rat [387], and GnRH and GAP have been found to be cosecreted into the hypothalamic–pituitary portal system in sheep [388]. The gene and hypothalamic cDNA for the common precursor for GnRH and GAP were isolated from libraries derived from human and rat hypothalamic mRNA [389]. The GAP sequences in human and rat have 17 amino acid differences, of which eight are conservative in nature [389]. However, GAP had no PRL suppressing activity when tested against human prolactinomas cultured in vitro [390] and neither GAP nor fragments of GAP were able to inhibit PRL release from lactating or oophorectomized, hypothalamo–pituitary disconnected ewes [391]. At this point the physiologic significance of GAP as a PIF vis-à-vis DA is still not clear. Still less is known about its bioactivity in humans. Because of the inverse nature of its suppressing effect on PRL while having stimulating effects on gonadotropins, GAP remains an intriguing unknown. g -Aminobutyric Acid (GABA)

Schally et al. [392] initially demonstrated the inhibitory effect of GABA on PRL secretion in vivo and in vitro in rats, and Grandison and Guidotti demonstrated high-affinity

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GABA receptors on the pituitary [393]. A tuberoinfundibular GABAergic system has been described with perikarya located in the arcuate nucleus and nerve endings demonstrated in the median eminence [394]. GABA has also been demonstrated to be present in portal blood [395]. High concentrations (10-5 mol/L) of GABA and muscimol, a GABA receptor agonist, have been shown to inhibit PRL synthesis and release in vitro in rats but blockade of this inhibition with GABA receptor antagonists, such as picrotoxin and bicuculline, have given variable results [393,396,397]. Subsequent studies have shown GABA and muscimol to cause a decrease in lactotroph PRL mRNA levels with in vitro incubations [398]. Studies of the effects of GABA in vivo in rats have shown variable results, probably due to inactivation of GABA by GABA transaminase. When inhibitors of the catabolism of GABA are employed, GABA can be shown to decrease circulating PRL levels and lactotroph PRL mRNA levels [398]. Studies of the GABA system in humans have yielded conflicting results in studies of widely differing experimental designs. Bioactive metabolites of GABA, such as gammahydroxybutyric acid and gamma-amino-beta-hydroxy butyric acid, and muscimol cause modest increases in PRL levels when injected into humans [399–401]. However, GABA itself causes a modest decrease in PRL levels when given to humans for several days [402] and activation of the endogenous GABAergic system with sodium valproate causes a suppression in the PRL rise induced by mechanical breast stimulation in puerperal women [403]. The physiologic role of GABA, like that of GAP, remains to be fully elucidated in the human as well as various animal species.

PRFs TRH

Shortly after its initial isolation and characterization, TRH was demonstrated to cause a rapid release of PRL from rat pituitary cell cultures [404] and in humans after intravenous injection (Fig. 4.9) [405,406]. Release of PRL is biphasic, the initial peak being mediated by activation of intracellular phosphoinositide pathways with IP3 generation and mobilization of intracellular calcium causing release of stored hormone; the second, more sustained phase is mediated by influx of extracellular calcium through calcium channels, which causes sustained secretion and synthesis of new hormone [166,407]. However, Tashjian et al. [408] found the rise in intracellular calcium to precede the rise in intracellular IP3, so that other mediators of intracellular calcium mobilization may also be involved. TRH is also internalized within the lactotrophs [409] and can even be synthesized by lactotrophs [410] but the physiologic significance of this is not clear. A number of different experimental approaches have failed to clarify the physiologic role of TRH as a PRF. The smallest dose of TRH that releases TSH also releases PRL in humans [411]. Immunoneutralization of endogenous TRH

FIGURE 4.10. Blockade of the prolactin rise induced by ether stress in rats treated with anti-vasoactive intestinal polypeptide (VIP) antiserum or normal rabbit serum (NRS). From Abe et al. [316]

with TRH antisera causes a 50% suppression of basal PRL levels in rats in some studies [412] but not in others [413,414]. Neither did such immunization affect the PRL response to electrical stimulation of the paraventricular nucleus or suckling [414], and it delayed but did not affect the magnitude of the spontaneous surge of PRL on proestrus [415]. Active, chronic immunization of ewes with TRH conjugated to albumin resulted in only minimal decreases in PRL levels basally and after stimulation by stress, suckling and estrus although reductions in thyroid hormone levels were more marked [416]. In the mouse with targeted disruption of the TRH gene (TRH KO), mice became hypothyroid with elevated levels of TSH with reduced biological activity but had normal PRL levels [417], further casting doubt on the essential role of TRH in PRL regulation. Suckling causes an increase in hypothalamic and portal vessel TRH levels as well as a decrease in DA levels [336]. If TRH mediates the PRL response to suckling, even in part, it ought to be accompanied by an increase in TSH, unless there were a concomitant increase in somatostatin. Detailed evaluations of serum levels of PRL and TSH following suckling in rats showed elevations of TSH in some studies [418,419] but not in others [414,420]. Similar studies in humans failed to show any elevations of TSH [421,422]. Very small doses of TRH given systemically were effective in releasing PRL and TSH in lactating rats and women in those studies, however, so it is unlikely that failure to show a rise in TSH was due to an increase in somatostatin. In hypothyroidism, TRH synthesis is increased [423] and portal vessel TRH levels are increased but not statistically significantly [424]. In addition in hypothyroidism, pituitary TRH binding is increased due to an increased number of TRH receptors, but an effect on binding affinity is contro-

Chapter 4

versial [425–427]. In human hypothyroidism, basal TSH and PRL levels are increased as are their responses to injected TRH [428]. Correction of the hypothyroidism with thyroid hormone corrects both the elevated TSH and PRL levels and their responses to TRH, although the correction is more rapid for TSH [428–430]. Conversely, in hyperthyroidism in humans, PRL levels are not low basally but the PRL response to TRH is markedly blunted and returns to normal with correction of the hyperthyroidism [428–430]. The above conflicting data from passive immunization studies, TRH KO mice, observation of TSH levels during lactation and examination of PRL levels in various thyroid states support a role for TRH as a physiologic PRF, albeit not the primary one or even one of major importance. TRH is itself generated from a larger precursor with a molecular weight of 29.2-kDa [431] and other peptides generated from the precursor have been found in the brain, but their functions are unknown [432]. Although also found in other nuclei, the neuronal perikarya for TRH which project ventrally to terminate in the median eminence are located in the parvocellular division of the paraventricular nucleus [431–433]. A stable, bioactive metabolite of TRH, cyclo-His-Pro (also known as histidyl-proline-diketopiperazine), is found in rat hypothalamus and pituitary in concentrations similar to those of TRH [434]. Cyclo-His-Pro inhibits the PRL response to TRH from rat pituitaries in vitro but reports conflict as to whether it inhibits the PRL response to TRH or suckling in vivo [435–438]. In conscious monkeys it also suppresses basal and TRH stimulated PRL levels [439]. The mechanism by which cyclo-His-Pro suppresses PRL release is not clear, as it does not appear to act via the TRH receptor [436]. How cyclo-His-Pro integrates with other factors involved in the regulation of PRL in animals and humans is as yet unknown. VIP and Peptide Histidine Methionine (PHM)/PHI

VIP was initially found to stimulate PRL release in 1978 [440,441]. VIP neuronal perikarya are present in the parvocellular region of the paraventricular nucleus, with axons terminating in the external zone of the median eminence [442,443]. Its effects are selective for PRL and additive to TRH in causing PRL release [444,445] at concentrations (2–6 ¥ 10-10 M) found in hypothalamic–pituitary portal blood [446,447]. As discussed above, the effects of VIP appear to be mediated by stimulation of adenyl cyclase, although recent evidence suggests that transport of calcium through membrane calcium channels may also be important [448]. In addition to stimulating PRL release,VIP also stimulates pituitary PRL mRNA content and PRL synthesis [449]. In conditions of increased PRL synthesis, such as lactation, hypothalamic VIP mRNA levels are also increased [450]. In addition to these studies in rats, intravenously administered VIP has also been shown to increase PRL levels in monkeys [451] and humans [452–454] at serum levels similar to those demonstrated in rat portal blood.

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FIGURE 4.11. Parallel fall in prolactin and growth hormone (GH) levels following resection of a growth hormone-releasing hormone producing pancreatic tumor that had caused acromegaly. From Thorner et al. [540]

A number of experiments have been performed using passive immunoneutralization techniques to determine the physiologic role of VIP as a PRF. Anti-VIP antisera administered to rats have been shown to partially inhibit the PRL responses to suckling, ether-induced stress (Fig. 4.10) [316,317], estrogen administration [455], intraventricular 5HT [456] and proestrus [457] and to suppress PRL pulsatile secretion in dopamine receptor blocked rats [458]. Similar results for suckling have been obtained with a VIP antagonist [459]. Part of the 20-kDa 170 amino acid VIP precursor is another similarly sized peptide known as PHM [460]. VIP has 28 and PHM has 27 amino acid residues. The terminal amino acid in the porcine equivalent of PHM is isoleucine, and therefore the peptide of porcine origin is known as PHI. PHI and VIP [461] and PHM and VIP [462] colocalize in the hypothalamus and median eminence. Interestingly, PHI is of similar potency to VIP in releasing PRL from rat pituitaries in vitro and in vivo [463,464]. Furthermore, passive immunoneutralization with anti-PHI plus anti-VIP antisera causes a greater suppression of the PRL responses to the serotonin precursor 5-hydroxytryptophan [465] and ether [317] than either anti-PHI or anti-VIP antisera alone. PHM given to humans has caused a PRL increment in some experiments [462,466] and not others [454]. Further complicating the role of VIP as a PRF is the finding that VIP is actually synthesized by anterior pituitary

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tissue [467]. Antisera to VIP inhibit basal PRL secretion from dispersed pituitary cells in vitro [468,469], suggesting a local “autocrine” role for VIP in PRL regulation within the pituitary. Interestingly, pituitary VIP concentration is increased in the pituitaries of hypothyroid rats along with a decreased PRL content [470] and TRH increased VIP and PRL secretion. However, immunoneutralization of the pituitary cultures with anti-VIP antiserum did not decrease the PRL response to TRH [471]. It is not yet clear whether the VIP secreting pituitary cells are lactotrophs [469] or nonlactotroph stellate cells [470]. The physiologic role of VIP as a PRF appears to be warranted by the experimental data. The precise roles of VIP vs. PHM and hypothalamic VIP vs. pituitary VIP still are not clear. How VIP/PHM interact with other PRF’s such as TRH are additional areas requiring clarification. Serotonin

A considerable number of experiments have demonstrated a role for serotonin as a neurotransmitter involved in the release of PRL. Most serotoninergic neuronal perikarya are in the dorsal and median raphe nuclei, and their axons project forward to the hypothalamus and other limbic and cortical areas [472]. Lesions of the dorsal but not the median raphe nuclei decrease forebrain serotonin levels and basal and stimulated serum PRL levels [472–475], while stimulation of the dorsal raphe nucleus results in PRL release [473]. Serotonin and its precursor, 5-hydroxytryptophan, cause a release of PRL in rats whether injected systemically [476,477] or into the third ventricle [478]. In a variety of experiments using blockers of serotonin synthesis, such as parachlorophenylalanine, blockers of serotonin receptors, such as methysergide, or blockers of serotonin reuptake by nerve terminals, such as fluoxetine, it has been shown that serotonin mediates, in part, the PRL elevations associated with suckling and proestrus [473,475,477,479–481]. Conversely, agents which are serotonin releasers, such as fenfluramine, or agonists, such as quipazine, result in a rapid increase in serum PRL levels [481–483]. Using specific serotonin receptor blockers, it has been found that it is the S2 receptor that mediates the serotonin action on PRL secretion [483]. Studies in humans also suggest a role for serotonin in PRL secretion. Infusion of the serotonin precursor, 5hydroxytryptophan, results in a prompt increase in PRL levels [484,485]. Nocturnal PRL secretion is inhibited by cyproheptadine [486]. Fenfluramine, a serotonin releasing agent caused a fourfold rise in PRL in humans that could be partially blocked by cyproheptadine [487]. Similarly, mchlorophenylpiperazine, a metabolite of the antidepressant drug, trazadone, has agonist activity at central serotonin receptors and causes a rise in PRL levels [488]. Fluoxetine, a serotonin reuptake inhibitor, also increases PRL levels modestly but still within the normal range [489]. On the other hand, pizotifen, a specific, non-ergot serotonin antag-

onist, had no effect on the suckling-induced PRL rise in postpartum women [490]. Whether serotonin’s effects are mediated solely through brain pathways or whether it has direct effects on the pituitary is controversial. One possibility is that serotonin causes a decrease in hypothalamic DA generation. Synaptic junctions between serotoninergic nerve terminals and dopaminergic perikarya in the arcuate nucleus have been demonstrated [491]. Furthermore, Pilotte and Porter [492] showed that intraventricular injections of serotonin decreased portal vessel DA concentrations and Johnston and NegroVillar [493] demonstrated that 5-hydroxytryptophan administration to rats results in a decrease in median eminence DA synthesis. However, when DA levels were maintained by infusion, serotonin was still able to cause a release of PRL [492]. It has also been proposed that serotonin acts by increasing VIP. Shimatsu et al. demonstrated that serotonin stimulates the release of VIP into portal blood [447] and passive immunization against VIP (see above) decreases the increase in PRL induced by 5-hydroxytryptophan [465]. Electrolytic lesions which destroy the paraventricular nucleus, the site of origin of VIPergic tuberinfundibular pathways, result in a marked blunting of the PRL responses to 5-hydroxytryptophan, ether stress and restraint stress [494], further buttressing the argument for a role for VIP in mediating the serotonin stimulatory effects on PRL secretion. Serotonin [495,496] and serotoninergic nerve fibers have been found within the anterior pituitary [497,498]. Serotoninergic nerve terminals have also been demonstrated within the median eminence [499]. Tryptophan hydroxylase, the rate-limiting enzyme in serotonin synthesis is present in the pituitary [496], suggesting in situ synthesis. High affinity S2 serotonin receptors have been found in the anterior pituitary [500] as well uptake of labeled serotonin into cells of the pituitary [501]. In direct tests of the effects of serotonin on pituitary PRL release, Apfelbaum [502] found serotonin to increase basal and stimulated PRL secretion from pituitaries in vitro and Stobie and Shin [503] and Wehrenberg et al. [504] found that serotonin stimulated PRL from hypophysectomized pituitary grafted rats and stalk-section monkeys, respectively. On the other hand, Birge et al. [329], Lamberts and MacLeod [505] and Delitala et al. [506] found no effect of serotonin on PRL release from rat pituitary cells in vitro. Thus, although it is possible that serotonin is a direct secretagogue for PRL, via transport from the hypothalamus by the portal vessels or through an autocrine action within the pituitary, its role in this regard is still uncertain. It likely mediates the nocturnal surge of PRL, and may well participate in the suckling-induced rise in PRL via the ascending serotoninergic pathways from the dorsal raphe nucleus and mediated by activation of VIP release. Opioid Peptides

A detailed description of the various opioid peptides, their receptors and their neuronal pathways is beyond the scope

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of this discussion. Approaches to determining the roles of the opioid peptides and pathways in the regulation of PRL secretion have focused on using opioid agonists and antagonists in experimental animals and humans. In rats, morphine, met- and leu-enkephalin, bendorphin, dynorphin and leumorphin injected systemically or intracerebroventricularly have all been shown to cause release of PRL [507–518]. More recent studies employing specific agonists and antagonists operative on the m, d and k opioid receptors and antibodies directed against several opioid peptides have shown that it is the m receptor that is the predominant one involved in PRL release, the k receptor is involved to a lesser extent and the d receptor is not at all involved [518–521]. Most evidence suggests that the opioid peptides do not have a direct effect on the pituitary [508,521,522] and stimulate PRL release by inhibiting DA turnover and release by the TIDA pathway [523–526]. The recently identified opioid peptide, orphanin FQ (also called nociceptin), which binds to an opioid-like orphan receptor, also causes an increase in PRL levels in rats when administered intracerebroventricularly [527]. In humans, morphine and morphine analogs increase PRL release acutely [528–532] and chronically [522,523]. However, blockade of the m receptor with naloxone has minimal to no effect on PRL levels either basally or with stimulation by hypoglycemia, exercise, sleep, TRH or physical stress [315,533–537]. In contrast to these findings, two groups have reported an increase in PRL levels in response to naloxone given in the late follicular and midluteal phases of the menstrual cycle [538,539]. The interpretation of these changes is not clear. Overall, it appears that the endogenous opioid pathways play at most only a minor role in the regulation of PRL secretion, especially in humans. GHRH

A number of studies have found GHRH to have PRL releasing properties. The initial clue to this effect of GHRH was the finding that many of the patients with acromegaly due to GHRH secreting tumors were hyperprolactinemic and PRL levels fell in parallel with GH following excision of the GHRH-secreting tumor [540] (Fig. 4.11). Large doses of GHRH have been reported to release PRL in vivo in normal humans in most studies [541–546]. In our own studies, GHRH was found to cause small, consistent PRL responses in addition to larger GH responses in normal individuals (Fig. 4.12) [543]. Chronic therapy with GHRH in children with GH neurosecretory dysfunction results in a sustained elevation of PRL levels [547]. In rat pituitary cell cultures, GHRH causes PRL and GH release but an increase only in GH mRNA and not PRL mRNA, indicating no stimulation of PRL synthesis [548]. The similarity of GH and PRL responses to a variety of stimuli, such as exercise, stress, hypoglycemia, arginine infusion and sleep and the pathological conditions of renal failure and hepatic cirrhosis suggest, but do not prove, that GHRH may serve as a physiologic PRF under some circumstances.

FIGURE 4.12. Prolactin secretory responses to bolus injection of growth hormone-releasing hormone (GHRH1–40) 3.3 mg/kg. Modified from Goldman et al. [543]

Posterior Pituitary, Oxytocin and Vasopressin

A number of studies in animals have shown that oxytocin, in levels found in the hypothalamic–pituitary portal vessels [549], can stimulate PRL release when added to the medium of pituitary cell cultures or incubations or when given intravenously but it lowers PRL levels when directly injected into the third ventricle [550–554]. Studies in which endogenous oxytocin was eliminated by passive immunization with oxytocin antisera or by oxytocin antagonists show a reduction and a delay in the suckling induced PRL surges in some studies [551] but not others [552] and a reduction in the PRL rise associated with proestrus [552,555] but no effect on the PRL response to ether stress [552]. Antisera to oxytocin also block the PRL surge seen in response to intracerebroventricularly administered VIP [459]. In addition, the peak phase of the PRL surge accompanying proestrus can be blocked by either posterior pituitary

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lobectomy or antisera to VIP [457]. These last findings suggest that oxytocin and/or other posterior pituitary substances may be involved in at least one of what are turning out to be multiple routes by which VIP stimulates PRL release. Very limited studies in humans suggest that oxytocin administered intravenously has no effect on basal PRL levels and causes only a minimal increase in TRH stimulated PRL levels [556]. Vasopressin also has PRL releasing properties when injected intravenously into normal rats and sheep and rats with pituitaries transplanted to the renal capsule, but not sheep with hypothalamic–pituitary disconnection [553]. The neurophysin portions of the precursors to oxytocin and vasopressin also stimulate PRL secretion in rats [557]. Schally et al. [558] have shown that a peptide identical to the sequence of 109–147 of the vasopressin–neurophysin precursor has been isolated from porcine hypothalami and has PRL inhibiting properties. Nagy et al. [559] found that the suckling induced PRL rise is absent in Brattleboro rats (which have diabetes insipidus due to a single base deletion in the vasopressin gene) and is partially restored with exogenous vasopressin. There are no studies to date of the effects of vasopressin on PRL secretion in humans. Whether there are other PRF’s in the posterior pituitary in addition to oxytocin, vasopressin and their respective neurophysins has been a matter of controversy. Rats which have had the posterior pituitary removed and replaced with vasopressin and oxytocin have a PRL response to suckling that is greatly reduced [560]. Antagonists to TRH, angiotensin II, endorphin and inactivation of vasopressin and oxytocin by disrupting disulfide bonds do not affect this posterior pituitary PRF activity [560,561] and studies of its physical characteristics suggest that it has a molecular mass of 1–5-kDa [562]. Although Mori et al. [554] could attribute posterior pituitary PRF activity only to contained oxytocin, Samson and Mogg [563] agree with Ben-Jonathan and her coworkers that the posterior pituitary PRF activity is due to some additional factor, and recently Laudon et al. [564] have localized the PRF activity to the intermediate rather than the true posterior lobe. The posterior pituitary, therefore, may play a significant role in the regulation of PRL secretion in rats. Vasopressin, oxytocin, the neurophysins and possibly an additional factor may be involved, possibly including VIP in the concatenation of neural links. How these factors interact with dopamine, TRH and other release and inhibiting substances in PRL regulation and how relevant all of this is to human PRL regulation are aspects that remain to be determined. Gonadotropin Releasing Hormone

GnRH was initially found to release PRL from rat pituitary cells in vitro [565]. Subsequently, GnRH has been found to cause a release of PRL in anovulatory women [566–568] and in women with anorexia nervosa who were gaining weight [569]. In 24% to 78% of normal women there is a PRL response to GnRH, depending upon the phase of the

menstrual cycle, the highest number of responders being in the periovulatory phase [570]. Postmenopausal women also have a PRL response to GnRH that is augmented with estrogen supplementation [571]. There is no PRL release in response to GnRH in normal, eugonadal males but such a release does occur with high doses of estrogen pretreatment (given to transsexual men) [572]. Analysis of PRL and LH secretory pulses suggests a high degree of concordance in rats and women [295,573]. This cosecretion of LH and PRL suggests that the response to GnRH is physiologic and is evidence against the inhibitory cosecreted GAP having a physiologic role. A subset of human prolactinomas that also contain the glycoprotein a subunit have been shown to bind GnRH specifically and with high affinity and to release PRL in response to GnRH in vitro [574]. Renin–angiotensin System

Angiotensin converting enzyme (ACE) and angiotensin II receptors and activity have been identified in the rat pituitary and median eminence of the hypothalamus [575–577]. In the human pituitary, renin, ACE and angiotensinogen have been detected in normal lactotroph cells and in PRLsecreting adenomas [578]. Angiotensin II incubated with rat pituitary cells stimulates release of PRL, an effect blocked by AT1 but not AT2 antagonists [579,580a]. However, in humans, blockade of ACE with enalapril results in no change in basal PRL levels [581,582a], no change in the TRH and metoclopramide induced PRL rises [581] and only a minimal decrease in the PRL response to hypoglycemia [582a]. It is unlikely, therefore, that the endogenous reninangiotensin system of the hypothalamus and pituitary have significant physiologic effects on PRL regulation. PRL-releasing Peptide

Hinuma et al. [582b] recently discovered a new, 31 amino acid peptide capable of releasing PRL, termed PrRP31, by looking for endogenous ligands for an orphan receptor present in the human pituitary termed hGR3. In pituitary cell preparations, PrRP31 released PRL with a potency equal to that of TRH [582b]. However, although PrRP31 is found in neuronal perikarya in the paraventricular and supraoptic nuclei, PrRP31-immunoreactive nerve fibers are found only in the internal zone of the median eminence and not the external zone [582c], casting uncertainty on the physiologic significance of this new peptide. Other Neuroactive Peptides and Neurotransmitters

Neurotensin This has been found to increase PRL levels when given intravenously to urethane-anesthetized intact and steroid primed male rats [508,583]. In in vivo studies, diphenhydramine, an H1 histamine receptor blocker, was able to block this stimulatory effect of neurotensin [508] but other studies have found neurotensin to release PRL directly from rat hemipituitaries in vitro [584]. When neurotensin is injected intracerebroventricularly, however, it inhibits PRL release [583,585] and the intracerebroventricular injection of

Chapter 4

antisera to neurotensin results in a rise in PRL levels, suggesting that endogenous neurotensin may be playing a physiologic role in rats [586]. In contrast to these results in rats, the administration of neurotensin to humans has no effect on PRL levels [587]. Substance P This also stimulates PRL release when given intravenously to urethane-anesthetized rats [508,588] and when given intracerebroventricularly to monkeys [589]. No such inhibition occurred when Substance P was given intracerebroventricularly to rats [585]; indeed, inhibition of PRL release was reported when it is given in this fashion [590]. Substance P also stimulates PRL release directly in vitro with pituitaries from female [585] but not male [590] rats. Cholecystokinin (CCK) This has been shown to increase PRL release from in vitro rat preparations [445,591] but not from similar human pituitary cultures [445]. The intravenous route in rats has provided both positive [591] and negative results [592] with respect to PRL stimulation. Intracerebroventricular CCK results in a marked PRL release that can be antagonized with antisera to VIP [592], implying that its action may require the activation of VIPergic systems. The intracerebroventricular or intravenous administration of proglumide, a CCK antagonist, cause a marked fall in PRL levels in male but not female rats [593]. The reason for these varying results, the discrepancy in findings between the sexes and the relevance to humans are unknown. Somatostatin Receptors for somatostatin have been found on human PRL as well as GH-secreting adenomas [594]. Somatostatin has been found to inhibit adenyl cyclase activity of rat anterior pituitary homogenates and spontaneous [595,596], TRH-induced [596], and VIP-induced [597] PRL release. Furthermore, administration of somatostatin antiserum to rats causes a rise in PRL levels, implying a physiologic inhibitory action of somatostatin basally [594]. In humans, however, somatostatin administered exogenously has no effect on the TRH-induced PRL release [598]. Other Peptides Bombesin is present in the hypothalamus and stimulates PRL secretion in vivo from steroid-primed male rats [599] and in vitro from pituitary cell cultures [600,601]. However, when bombesin is administered intracerebroventricularly, it inhibits the PRL rise induced by restraint stress [602]. No studies of its effects in humans have been reported. Secretin has been found to have both stimulating [603] and inhibiting [604] effects on PRL release in rats. Gastrin causes an increase in PRL levels when injected intravenously in rats but causes a fall in PRL levels when injected intracerebroventricularly. It has no effect on PRL secretion in vitro [605]. Galanin is expressed in the PVN and the median eminence of the hypothalamus and in pituitary lactotrophs, where it is colocalized with PRL in the same secretory granules [606]. Galanin injected intravenously

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causes an increase in PRL levels when given in high doses, but has no effect when injected intracerebroventricularly or in vitro [607]. Galanin KO mice have 30% to 40% lower PRL levels, are unable to lactate, and their lactotrophs fail to proliferate in response to estrogen [606]. Endothelin is produced by lactotrophs and also causes a release of PRL from lactotrophs, suggesting a possible autocrine regulation of PRL secretion [608]. Transforming growth factor-b1 (TGF-b1) is similarly produced by lactotrophs but inhibits PRL gene expression, synthesis and release in a paracrine rather than an autocrine manner [609,610]. Adenosine is produced by the folliculostellate cells of the pituitary and also stimulates PRL release in a paracrine fashion [611]. Several other substances have been found to increase PRL release from pituitary cell cultures, including bradykinin [612,613], calcitonin [614]; calcitonin generelated peptide (CGRP) [615]; relaxin [616], plateletactivating factor [617], and thymosin fraction 5 [618]. Melatonin has also been shown to increase PRL levels from humans when given orally [619]. On the other hand amelanocyte stimulating hormone (a-MSH), injected intracerebroventricularly causes a decrease in PRL levels and the administration of MSH antiserum causes a rise in PRL levels [620,621]. Basic fibroblast growth factor also inhibits PRL release in vitro [622]. A peptide identical to the 27–52 amino acid sequence of proopiomelanocortin has been isolated from the hypothalamus and is also inhibitory to PRL secretion [558]. Pituitary adenylate cyclase activating polypeptide (PACAP) also inhibits PRL secretion when injected intracerebroventricularly through a stimulation of DA release by TIDA neurons [623]. Histamine This neurotransmitter plays an uncertain role in PRL regulation [624]. Histamine neuronal perikarya are present in the posterior hypothalamic region and axons project to almost all of the nuclei of the hypothalamus [624]. Histamine administered systemically or intracerebroventricularly to rats causes a modest PRL rise [625–627]. H1 receptor antagonists suppress the suckling-induced PRL increment in rats [625] as well as the PRL rise caused by intravenous histamine [627], whereas H2 receptor antagonists, such as cimetidine and ranitidine, augment the intravenous histamine-induced PRL rise [628]. In contrast, intracerebroventricular histamine also increases PRL release, an effect blocked by H2 blockers but not H1 blockers [628]. These studies suggest that histamine exerts its PRLreleasing activity via H2 receptors when given intracerebroventricularly and via H1 receptors when given intravenously. However, histamine has no effect on PRL release from pituitaries in vitro or in stalk-sectioned rats [627]. Although intracerebroventricular histamine causes a 26% fall in stalk vessel dopamine levels, this is not sufficient to explain the large rise in plasma PRL levels [626]. In humans, intravenous H2 but not H1 blockers cause a rise in PRL levels [629–632] but prolonged oral administration of H2 blockers does not result in sustained PRL elevation

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[632]. Histamine alone can cause a rise in PRL secretion in man that is further augmented by H2 blockers and is blocked by H1 blockers [628]. The fact that the administration of high doses of H2 blockers causes an increase of PRL levels in humans, that histamine cannot cross the blood–brain barrier and that histamine has no effect on pituitaries in vitro suggests that histamine may play a physiologic facilitory role within the median eminence in PRL secretion. The roles of other bioamines in the regulation of PRL secretion are even less well established. Central adrenergic a2 agonists such as clonidine usually have no effect on PRL secretion although both increases and decreases have been reported, the differences possibly being due to the doses used [633–635]. The administration of a2 adrenergic antagonists such as yohimbine or piperoxan result in an increase in PRL levels that is blocked by clonidine [635,636], implying an overall inhibitory role of the a2 adrenergic system in the hypothalamus. a-methyl dopa, another central adrenergic agonist, causes a sustained elevation of PRL [637] but this may be due to inhibition of the synthesis of norepinephrine or dopamine centrally by inhibiting the enzyme l-aromatic acid decarboxylase, which is responsible for conversion of dopa to dopamine and by acting as a false neurotransmitter to decrease dopamine secretion or synthesis by a local feedback inhibitory action. However, monoamine oxidase inhibitors cause an increase in PRL levels [638]. Such an elevation is unexpected, since these drugs should increase the levels of norepinephrine and dopamine in the synapse. Acetylcholine This neurotransmitter inhibits adenyl cyclase and cAMP accumulation, lowers intracellular free calcium levels and decreases PRL release from pituitary cell cultures, acting through muscarinic and not nicotinic receptors [127,639]. In such cultures, coincubations of acetylcholine or the cholinergic agonist, carbachol, with VIP or TRH result in marked inhibition of the VIP- and TRHinduced PRL release [127,640]. Although atropine, a muscarinic receptor antagonist, blocks this acetylcholine inhibition of PRL in vitro [127,640], it had no effect on basal or TRH-induced PRL release in humans [641]. Moreover, pirenzepine, another muscarinic receptor blocker, actually caused a modest decrease of PRL levels in humans in vivo [642]. Another muscarinic receptor blocker, methscopolamine, had no effect on the sleep induced PRL rise while blocking the GH rise [643]. The widespread presence of acetylcholine as a neurotransmitter in the CNS and the possibility that pituitary tissue itself may synthesize acetylcholine [644] make interpretation of studies testing this system difficult, and the true role of acetylcholine in the regulation of PRL secretion is uncertain.

Prolactin Short Loop Feedback Considerable evidence in rats suggests that PRL is able to feed back negatively on its own secretion (short-loop feedback or autofeedback) [645–647]. Most evidence

suggests that such feedback occurs via augmentation of hypothalamic TIDA turnover [648–652], including direct measurements of dopamine in portal vessels [379]. Direct confirmation of the importance of short-loop feedback in rodents comes from recent studies using mice with targeted disruption of the PRL gene (PRL gene KO). Mice with the PRL-KO have no pituitary PRL and have markedly decreased DA in TIDA neurons along with hyperplasia of lactotrophs that do not make PRL [653–655]. Direct evidence for such PRL short-loop feedback in the human has not been demonstrated. In a number of reports, however, it has been suggested that altered regulation of gonadotropin and TSH secretion in hyperprolactinemic patients may constitute indirect evidence of PRL-induced augmented TIDA activity [656–663]. In hyperprolactinemic patients, decreases in gonadotropin pulse amplitude and frequency are usually found, being attributed to altered gonadotropin releasing hormone secretion (see below). Such an alteration of GnRH secretion has been postulated to be due, in part, to PRL-induced DA increase [656,664–667]. In a direct test of this hypothesis, however, no evidence was found of suppression of TSH, LH or FSH levels with short term administration of human PRL which resulted in a two- to threefold elevation of PRL levels [668]. However, these acute studies do not rule out the possibility that such feedback may occur with more prolonged states of hyperprolactinemia. Alternatively, such feedback might occur via other mechanisms, such as a decrease in a PRF such as VIP [669]. PRL ACTION PRL has a great diversity of actions in many species of animals from fish and birds to mammals, including osmoregulation, growth and developmental effects, metabolic effects, actions on ectodermal and integumentary structures and actions related to reproduction [670]. However, in humans it has as its primary physiologic action the preparation of the breast for lactation in the postpartum period [671]. A number of effects of increased levels of PRL may be seen on many tissues. Although the roles of physiologic levels of PRL in such tissues are quite speculative, considerable clarification of these roles has been elucidated from studies in PRL-KO and PRL receptor KO mice.

PRL Receptors The PRL and GH receptors are members of the class 1 cytokine receptor family. The human genes for the PRL and GH receptors have been localized to the short arm of chromosome 5, sublocalizing to region 5p13–p14 [672]. The human PRL receptor gene has 10 exons, with exons 3 to 10 encoding the full length of the long form of the receptor [673]. Two PRL receptor gene promoters, termed hPIII and hPN that direct alternative first exons of the HPRL receptor gene have been found in the 5¢ untranslated region [673]. The human PRL receptor is 598 amino acids long,

Chapter 4

has a theoretical, non-glycosylated molecular mass of 66.9kDa and an additional signal peptide of 24 amino acids [674]. The extracellular domain contains five cysteine residues and three potential N-linked glycosylation sites [674]. There is a hydrophobic region (amino acids 211–234) which corresponds to the single transmembrane-spanning region of the receptor [674]. The human PRL receptor has a much longer intracytoplasmic region than does the rat PRL receptor [63] but there is high sequence homology between the rat, rabbit and human PRL receptors and also between the human PRL and GH receptors [63]. Two isoforms of the PRL receptor result from alternative splicing and differ in the length and composition of the cytoplasmic tail, being referred to as the long and intermediate forms; the short form found in the mouse is not present in humans [63,633]. PRL binds to its receptor with high affinity, the dissociation constant (Kd) being 10-10 mol/L [63]. Half-saturation of the receptor occurs at a hormone concentration of 7 ng/ml [63]. Once PRL binds to the receptor there is dimerization of the receptor, a necessary step for activation of the receptor (Fig. 4.13) [63]. PRL receptors are widely distributed, being found in the breast, pituitary, liver, kidney tubules, adrenal cortex, prostate, ovary, testes, seminal vesicles, epididymis, intestine, skin, pancreatic islets, lymphocytes, lung, myocardium and brain [63]. PRL release caused by suckling increases PRL receptor levels in the breast and liver, resulting in a much greater PRL binding activity in lactating animals compared to those not lactating [63]. Hepatic PRL receptor synthesis and numbers increase with estrogen treatment in vivo, mediated by the effect of estrogen on increasing pituitary PRL secretion [675] (Table 4.3). Interaction of PRL binding site 1, encompassing several amino acids belonging to helices one and four, with one PRL receptor causes the formation of a one-hormone-onereceptor complex. This is a prerequisite for PRL binding site 2, involving helices 1 and 3, to interact with another receptor, ultimately causing the formation of the active trimeric complex [63]. Signal transduction of the activated receptor involves the JAK-Stat pathway. JAK2 is the particular Janus kinase involved and it is constitutively associated with the PRL receptor [63]. PRL-induced dimerization of the two receptor molecules brings the two associated JAK2 molecules close to each other so that they may be activated by transphosphorylation of tyrosines [63]. The two cytoplasmic domains of the receptor must be strictly identical, and the short receptor functions as a dominant negative isoform which inhibits activation of the receptor complex by heterodimerization [63]. The activated JAK2 phophorylates tyrosine residues on the receptor itself and on three members of the Stat (signal transducer and activator of transcription) family of proteins, Stat 1, Stat 3 and, particularly, Stat 5 [63]. Activation of the MAP kinase cascade has also been reported after receptor activation but whether this involves JAK2 is not known [63]. Subsequently, the hormone–receptor complex is internalized and localized in the golgi and vacuoles [676] but the physiologic

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FIGURE 4.13. (a) Ribbon representation of the predicted 3D structure of hPRL, modeled on the basis of the crystallographic structure of porcine GH. HPRL is predicted to adopt the four-helix bundle folding described for GH. Location of binding sites 1 and 2 is indicated. Side chains of amino acids involved in binding site 1, as deduced from mutational studies are represented. (b) Human prolactin receptor extracellular domain. The extracellular domain folds in a b-sandwich formed by two antiparallel b-sheets and Cterminal ends are indicated by N and C respectively. Note that the structures depicted in panels A and B are not at the same scale. (c) Prolactin receptor activation induced by PRL-induced dimerization. Hormone binding to PRL receptor is sequential. First, the hormone (H) interacts with the receptor (R) through its binding site 1 (see A), forming an inactive H1 : R1 complex. Then, the hormone binds to a second receptor through its site 2, which leads to receptor homodimerization and formation of an active H1 : R1 complex. Hormone analogs whose binding site 2 is sterically blocked are unable to induce receptor homodimerization and are thus inactive; since they still bind to the receptor through site 1, they behave as antagonists of wild-type hormones. From Bole-Feysot et al. [63]

significance of this beyond degradation and scavenging is not known.

PRL Effects on Breast PRL, GH, cortisol, insulin, estrogen, progesterone and thyroxine all contribute to breast development. The high concentrations of estrogen and progesterone produced by the placenta, coupled with the estrogen-induced high concentration of circulating PRL and the high concentrations of hPL cause development of the lobular, alveolar tissue during

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Table 4.3.

Summary of phenotypes of PRL receptor +/- and receptor -/- mice*

+/-

-/-

Reproduction Female

Normal mating, pregnancy

Male

Normal

Sterility Ovulation decreased Oocyte maturation decreased No implantation Delayed fertility

Mammary gland development

Impaired development, efficient lactation after first pregnancy

No lactation Ductal branching greatly reduced Absence of alveoli

Behavior Maternal Eating, locomotor

Reduced Normal

Absent Normal

Immunity

Normal

Normal

* Modified from Bole-Feysot et al. [63]. +/-, heterozygous PRL receptor knockout, -/-, homozygous PRL receptor knockout.

pregnancy [671]. Once the breast is fully developed and hormonally primed, PRL stimulates the production of milk proteins and other components [671]. Prior to term, the high estrogen levels suppress the effects of the high PRL levels on milk production but the rapid fall off of estrogen levels following delivery allows milk production to proceed [677]. Suppression of this physiologic hyperprolactinemia in the puerperium by bromocriptine causes a rapid cessation of milk production [678]. The key role of PRL in milk formation has been demonstrated by the finding that mice with either the PRL gene KO [653] or the PRL receptor KO [679] are unable to lactate. One intriguing finding is the production by the lactating breast of the parathyroid hormonelike protein that has been implicated in the humoral hypercalcemia of some malignancies [680]. Suckling results in an increase in serum PRL and the appearance of the mRNA for this parathyroid hormone-like protein in breast tissue, both effects being blocked by bromocriptine treatment [680]. This protein is not present in the circulation in significant amounts but is present in breast milk in high concentrations [681], suggesting that it may act within the breast to facilitate calcium transport into the milk. Galactorrhea

Clinically, nonpuerperal galactorrhea has been regarded as being a sign of possible hyperprolactinemia. The presence of even minute amounts of milk expressible from one or both breasts justifies the diagnosis of galactorrhea. Its persistence for more than 1 year after normal delivery and cessation of breast-feeding or its occurrence in the absence of pregnancy generally is taken as a definition of inappropriate lactation. If the material expressible from the nipple looks like milk, it probably is milk; if there is any question

of diagnosis, examination of the breast secretion by staining of fat globules with Sudan IV is diagnostic. The incidence of galactorrhea has been variously reported in normal women as ranging from 1% to 45% of subjects tested [682–685]. This variability is probably the result of differences in the techniques used to express milk from the breast and the way in which nonmilky secretions are classified. The volume of milk expressed does not correlate with PRL levels. However, in individuals with hyperprolactinemia, lowering the blood PRL level to normal will almost always lead to a marked decrease or abolition of lactation. Inappropriate lactation may be an important clue to the presence of pituitary–hypothalamic disease, especially if accompanied by amenorrhea. In a series of 70 women with galactorrhea studied in the author’s clinic, 19 (27%) had normal menses [686]. Of those with normal menses, only one had elevated PRL levels. Combined data from 14 published series suggest that 27.9% of galactorrheic women with normal menses have elevated PRL levels [687]. More recent experience, however, suggests that galactorrhea may be present in about 5% to 10% of normally menstruating women and basal PRL levels are normal in more than 90% of these women.

PRL Effects on Gonadotropin Secretion The effects on gonadotropin secretion by PRL levels in the normal range are not known. However, PRL gene KO and PRL receptor KO female mice are sterile. Both types have disordered estrous cycles [653,679]. Females with the PRL receptor KO had fewer primary follicles, fewer eggs ovulated, fewer eggs fertilized, those that were fertilized had

Chapter 4

poor progression to the blastocyst stage and finally the uterus was refractory to implantation by the blastocyst [679]. Furthermore, they had decreased estradiol and progesterone levels [688]. Half of the males with PRL receptor mutations were fully fertile, but the remainder were either completely or partially infertile despite normal sexual behavior and normal spermatogenesis as determined by histologic evaluation of the testes [679]. Male mice with the PRL gene KO were fully fertile and had normal plasma testosterone levels and normal testosterone release from the testes, despite decreased plasma LH levels and LH and FSH secretion from the pituitary in vitro and decreased weights of the seminal vesicles and ventral prostate [689]. In normal women treated with short-term bromocriptine to lower PRL levels to about 5 ng/ml, there was no change in the pulsatile secretion of LH and FSH, but estradiol levels were higher the last 3 days of the follicular cycle and progesterone levels were lower during the luteal phase [690]. Hyperprolactinemia has a number of effects on various steps in the reproductive axis (Fig. 4.14). Hyperprolactinemia has been found in most studies to suppress LH pulsatile secretion by decreasing pulse amplitude and frequency [664,665,693–695]. With castration in rats or menopause in humans, hyperprolactinemia can prevent the expected rise in gonadotropins [696–699]; normalization of PRL levels with bromocriptine results in elevation of gonadotropin levels and hot flashes [699]. Hyperprolactinemia inhibits pulsatile gonadotropin secretion by a number of mechanisms. It had been postulated that the pulsatile gonadotropin secretion was directed by the hypothalamic GnRH pulse generator and that alteration of pulsatile secretion necessarily meant a direct hypothalamic action. However, direct measurement of portal vessel GnRH levels in rats showed a marked inhibitory effect of hyperprolactinemia in one study [700] but not in another [701]. The pituitary gonadotroph response to GnRH in hyperprolactinemia has generally been found to be decreased in rats [695,696,701] and normal, increased and decreased in humans [702–704]. In our own series, the gonadotropin response to GnRH was normal in 22 of 25 patients and decreased in the remaining three [686]. GnRH plays a major role in the regulation of the number of its own recep-

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tors on the gonadotroph cell. The number of GnRH receptors on gonadotroph cells in hyperprolactinemic rats are reduced [705] even when endogenous GnRH is replaced with intraarterial pulses of GnRH [695,706,707]. In addition to these effects, hyperprolactinemia in women has been associated with loss of positive estrogen feedback on gonadotropin secretion [708].

PRL Effects on the Ovary PRL has long been known to be trophic to corpus luteum function in rats, giving rise to the name luteotrophic hormone [709]. As noted above, the ovaries of PRL receptor KO mice have a decreased number of primary follicles with a decrease in the number of eggs ovulated [679] and decreased estradiol and progesterone production [688]. The role of PRL in normal ovarian function in humans is less well established, however. McNatty et al. [710] demonstrated that low physiologic concentrations of PRL are necessary for progesterone synthesis by human granulosa cells but that high concentrations are inhibitory in vitro. Recent studies suggest that PRL can activate the expression of Type II 3bhydroxysteroid dehydrogenase, the final enzymatic step in progesterone biosynthesis [711]. Del Pozo et al. [712] found no effect on luteal function in women treated with bromocriptine to lower normal PRL levels. However, Schulz et al. [713], Mühlenstedt et al. [714], and Kauppila et al. [690] found that such a decrease in PRL resulted in lowered progesterone levels and short luteal phases. On the other hand, short luteal phases have also been reported in hyperprolactinemic women [715]. In humans, plasma PRL levels > 100 ng/ml have been found to cause an increase of antral fluid PRL levels and reductions in antral fluid FSH and estradiol levels along with a decrease in granulosa cells [716]. Perfusion studies of human ovaries in vitro reveal a direct suppressive effect of PRL on progesterone and estrogen secretion [717]. PRL can inhibit estrogen formation by antagonizing the stimulatory effects of FSH on aromatase activity [718,719] and direct inhibition of aromatase synthesis has recently been shown [720]. PRL also inhibits LH-stimulated androgen synthesis by ovarian interstitial cells in rats [721], thereby

FIGURE 4.14. How hyperprolactinemia produces hypogonadism. Schematic representation of sites where hyperprolactinemia interferes with the reproductive system and sex steroid action. CNS, central nervous system; FSH, folliclestimulating hormone; LH, luteinizing hormone. From Odell [691], as modified by Malarkey [692]

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depriving the ovary of the substrate for conversion to estrogen. On the other hand, no differences in the amount of exogenous gonadotropins are necessary to induce ovulation [722,723]. PRL levels have been found to be elevated in 19% to 50% of women with polycystic ovaries (PCO) [724–728]. Bromocriptine treatment of hyperprolactinemic patients with PCO usually results in a reduction of testosterone and LH levels and resumption of ovulatory cycles. Why many patients with PCO have hyperprolactinemia is not clear. Del Pozo and Falaschi [726] have hypothesized that the increased estrogen levels found in PCO stimulate increased PRL secretion; however, no correlation has been found between estrone levels and PRL levels in these patients. The increased androgen secretion associated with PCO may cause hirsutism and other excessive androgen action such as acne and seborrhea [729], although these features may also be caused by excessive androgen secretion by the adrenal (see below). Clinical Effects of Hyperprolactinemia on Menstrual Function

When amenorrhea or oligoamenorrhea is associated with galactorrhea, it is usually a manifestation of hyperprolactinemia. In our series of 51 women with galactorrhea– oligo/amenorrhea, 84% had hyperprolactinemia [686]. In combined series totaling 471 patients with galactorrhea– amenorrhea, 75.4% were found to have hyperprolactinemia III [687]. Although the amenorrhea caused by hyperprolactinemia usually is secondary, it also can be primary if the disorder begins before the usual age of puberty. Of 33 patients presenting with primary amenorrhea and low gonadotropin levels, 9 (27%) were found to have hyperprolactinemia in two series [730,731]. However, in a third series of 38 patients with primary amenorrhea, PRL levels were normal in all [732]. In patients with primary amenorrhea due to hyperprolactinemia, estrogen deficiency and failure to develop normal secondary sexual characteristics may be the presenting problem. Galactorrhea is variable in this setting because the breast may not have been exposed to appropriate priming with estrogen and progesterone. Patients with primary amenorrhea tend to have macroadenomas more commonly than those with secondary amenorrhea, for uncertain reasons (see Prolactinoma, Chapter 12, p. 470). Hyperprolactinemia has been found in many women with a short luteal phase, as noted above. It is likely that a short luteal phase is the first evidence of interference in the normal cycle by hyperprolactinemia. Three hyperprolactinemic patients with amenorrhea corrected by bromocriptine studied by Seppala et al. [733] displayed the following sequence of events when bromocriptine was discontinued: (i) as the serum PRL levels increased, the luteal phase became shorter; (ii) ovulation was missed; and (iii) amenorrhea ensued. In the initial period of shortened luteal phase, progesterone levels were subnormal, suggesting deficient corpus luteum function.

Infertility also may be a presenting symptom of patients with hyperprolactinemia and is invariable when gonadotropin levels are suppressed with anovulation. In three series of women (combined number of 367 cases) studied for infertility, one-third were found to have hyperprolactinemia [687]. Most of the women had amenorrhea and galactorrhea as well, but in one series of 113 cases of infertility, five of the 22 hyperprolactinemic women had neither amenorrhea nor galactorrhea [734]. That PRL excess may be important in this type of patient is suggested by the finding that treatment of similar patients with bromocriptine restored fertility [729]. In some of these women, transient hyperprolactinemia lasting for 1–2 days during the cycle can be documented; this subset usually responds to bromocriptine with increased progesterone during the luteal phase and improved fertility [735]. Reduced libido and orgasmic dysfunction are found in most hyperprolactinemic amenorrheic women when such complaints are specifically elicited [686,736]. Reduction of PRL levels to normal restores normal libido and sexual function in most of these women [737].

PRL Effects on the Testes The role of PRL in normal testicular function is unclear. In hypophysectomized mice, Bartke [738] found that PRL increased the testicular spermatogenic response to LH and the administration of LH and PRL to such mice increases testosterone levels more than treatment with LH alone [739]. Suppression of normal PRL levels causes a decrease in binding of LH to LH receptors in Leydig cells [740]. In animals with hereditary PRL deficiency, PRL administration increases the activities of 17-beta-hydroxysteroid dehydrogenase and 3-beta-hydroxysteroid dehydrogenase, enzymes necessary for testosterone synthesis [741,742]. However, as noted above, the PRL gene KO mice had normal fertility and normal testosterone production despite decreased LH and FSH secretion [689], and half of the males with PRL receptor mutations were fully fertile with the remainder subfertile despite normal sexual behavior and normal spermatogenesis [679]. Suppression of normal PRL levels by bromocriptine in normal men for 8 weeks results in suppression of basal and hCG-stimulated testosterone levels, implying a physiologic role for PRL in normal testosterone production in humans [743]. Although PRL is present in human semen in very high concentrations [744] and PRL has been shown to stimulate adenyl cyclase, fructose utilization and glycolysis and glucose oxidation in human spermatozoa [745], other functions have not been investigated in detail. Chronic hyperprolactinemia in males results in impotence and decreased libido in over 90% of cases [746–754]. Other findings of hypogonadism, such as decreased beard growth and strength are less common [749,754]. Galactorrhea in men has been reported in 10% to 20% of cases and is virtually pathognomonic of a prolactinoma [746,749,751,754,755].

Chapter 4

In hyperprolactinemic men there is a decrease in the pulsatile secretion of LH and FSH, as noted above, and testosterone levels are low or are in the lower part of the normal range [746–754]. The testosterone response to stimulation with hCG has been reported to be both decreased [750,756] and normal [746,757]; in those with decreased responses there is improvement in the response when PRL levels are lowered with bromocriptine [750]. If there is sufficient normal pituitary tissue, reduction of elevated PRL levels to normal usually results in a return of normal testosterone levels [752,753,758,759]. Although some studies have suggested that drug-induced elevated PRL levels cause a partial block in the enzyme 5-alpha reductase, resulting in a decrease in dihydrotestosterone levels [760] this has not been found in studies of men with prolactinomas [756]. Carter et al. [746] noted that testosterone therapy of hyperprolactinemic men does not always correct the impotence until PRL levels are brought down to normal. Whether this is due to a decrease in DHT levels has not been verified directly. Sperm counts and motility are decreased with an increase in abnormal forms [757] and histological studies reveal abnormal seminiferous tubule walls and altered Sertoli cell ultrastructure [761]. The semen analysis does not always return to normal despite a return to normal of testosterone levels with restoration of normoprolactinemia [758]. A number of surveys have attempted to assess the frequency of hyperprolactinemia among men with complaints of impotence or infertility. Between 2% and 25% of males with impotence have been found to be hyperprolactinemic in various series [729,750,752,762–764]. However, only 1% to 5% of men with infertility have been found to be hyperprolactinemic [765–768]. Thus, although these frequencies are relatively low, the cost of the test is low in terms of money and risk and the condition is generally easily treatable, so that measurement of PRL levels is worthwhile in the investigation of the man complaining of infertility or impotence, especially if there is associated loss of libido or documented low testosterone levels.

Prolactin Effects on the Adrenal Cortex Although PRL receptors are found on cells of the adrenal cortex, the physiologic role of PRL in adrenal steroidogenesis is unknown. Plasma dehydroepiandrosterone (DHEA) and DHEA sulfate (DHEAS) levels have been found to be mildly elevated in about 50% of women with hyperprolactinemia in most [769–771] but not all series [772–774]. In most of these studies, however, the investigators did not try to correlate the androgen levels with the presence of hirsutism or other indices of virilization. Glickman et al. [774] and Vermuelen et al. [775] found low plasma sex hormone-binding globulin and elevated free testosterone levels in 43% of hyperprolactinemic patients, but increased DHEAS levels in only 19%. When the patients studied by Glickman et al. [774] were divided into those who were hirsute and those who were not, the hirsute patients had

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higher free testosterone levels but not higher DHEAS levels, although both groups had patients with elevated levels of both hormones. The abnormal androgen levels return to normal with correction of the hyperprolactinemia by bromocriptine [776].

Effect of Prolactin on Bones PRL may have a physiologic role in calcium and bone metabolism. PRL has been shown to increase intestinal calcium absorption even in vitamin D deficient rats [777] and can stimulate 1-a hydroxylation of 25 hydroxyvitamin D in the kidney, resulting in increased plasma levels of 1,25(OH)2D [778]. In humans, however, plasma 1,25(OH)2D levels and intestinal calcium absorption are normal in hyperprolactinemic subjects [779]. The PRL receptor KO mouse has a decrease in bone formation rate and bone mineral density in association with increased parathyroid hormone (PTH) levels, but also decreased estradiol and progesterone levels [688] so that it is difficult to say how much of the effect on bone is due to the lack of PRL. 1,25(OH)2 enhances PRL gene expression with dependence on extracellular calcium [780,781] and treatment of rats with 1,25(OH)2D causes an increase in serum PRL levels [782]. Recent studies suggest that this 1,25(OH)2D mediated increase in PRL is mediated by enhancement of calcium transport through membrane calcium channels [783]. The initial observation by Klibanski et al. that hyperprolactinemic women have a decreased bone mineral density [784] was confirmed by others [785,786] but whether this effect is mediated by estrogen deficiency [784] or is a direct effect of the hyperprolactinemia [785,787] has been controversial. Correction of the hyperprolactinemia results in an increase in bone mass [788,789]. Studies of hyperprolactinemic women who were not amenorrheic and hypoestrogenemic have shown that their bone mineral density is normal [790,791], confirming the initial hypothesis that it is the estrogen deficiency that mediates the bone mineral loss. A similar, androgen-dependent loss of bone mineral is found in hyperprolactinemic men that is reversible with reversal of the hypoandrogenic state [792,793].

Effect of Prolactin on Carbohydrate Metabolism PRL receptors have been found in islet cells [794] but the role of PRL in normal carbohydrate homeostasis is not known. Patients with hyperprolactinemia with or without evidence of prolactinoma have mildly increased (still within the normal range) blood glucose levels during glucose tolerance tests compared with normal controls [795,796]. Insulin levels are also increased in these patients, indicating mild insulin resistance [795,796]. The insulin resistance is due to a decrease in the number of receptors rather than a decrease in receptor affinity [797]. Treatment with bromocriptine causes a return to normal of both glucose and insulin levels [795]. However, in normoprolactinemic

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subjects with insulin-dependent diabetes, suppression of PRL levels with bromocriptine had no effect on glucose control [798].

Effects of Prolactin on the Kidney An extensive body of literature documents the importance of PRL in osmoregulation in fish, amphibians, reptiles, birds and lower mammals [799]. PRL effects include increased tubular reabsorption of sodium, chloride and water, increased glomerular filtration rate, increased thirst, and increased intestinal absorption of sodium, chloride, calcium, water, glucose and amino acids [799]. A number of experiments in humans, however, have failed to document a significant role for PRL in fluid and electrolyte balance or of osmotic stimuli for PRL release [800–803].

Effects of Prolactin on the Immune System Normal PRL levels appear to be essential for normal immune function in rats, although the precise role of PRL in humans in this regard is not clear. In hypophysectomized rats, antibody production against sheep red blood cells, the skin response to dinitrochlorobenzene and the rejection of skin grafts are all decreased [804]. These immune responses could be restored with treatment of the hypophysectomized rats with rPRL, rGH, bPRL, bGH, hPL and hGH [805]. Hypoprolactinemia induced with bromocriptine results in impaired lymphocyte proliferation and decreased production of macrophage activating factors by lymphocytes as well as decreased tumoricidal activation of macrophages after infections; these effects could be reversed with ovine PRL administration to the rats [806]. Neutralization of PRL with anti-PRL antibodies inhibits the in vitro lymphocyte proliferative response to T and B cell mitogens in both rat and human preparations, but neutralization with antibodies to other pituitary hormones had no effect [807]. However, when the lymphocytes were cultured in serum-free medium, the anti-PRL antibodies similarly inhibited lymphocyte proliferation, implying the presence of a necessary, lymphocyte-produced PRL-like protein [807]. In other studies, in response to stimulation with conconavalin A, a PRL-like substance has been found to be produced from murine splenic mononuclear cells that reacts with anti-PRL antibodies and is mitogenic in the Nb2 node lymphoma bioassay [808] and mRNA obtained from such lymphocytes hybridizes with PRL and GH cDNA probes [809]. Furthermore, the immunosuppressive drug cyclosporin A has been found to compete with PRL for a common binding site on rat lymphocytes and stimulation of PRL secretion reverses the immunosuppression induced by cyclosporin [809]. Presumably, this rat lymphocytic PRL-like substance is analogous to the lymphoblastoid PRL produced by IM9 lymphocytes discussed above [235]. In preliminary studies with the PRL gene KO and receptor KO mice, no effects were seen on B or T lymphocyte production but more

detailed studies on immune function have not yet been carried out [63,653]. Lymphoblastoid prolactin can also be produced by conconavalin A or phytohemag-glutinin stimulated human peripheral blood mononuclear cells and has been shown to cause lymphoproliferation, a phenomenon blocked by antihuman PRL antibodies [810]. Human T lymphocytes contain the genes for hPRL and the PRL receptor, but the lymphocyte hPRL gene has an extra 5¢ noncoding exon similar to that found for decidual PRL [811]. PRL mRNA has been localized to a variety of human immune tissues, including thymus, spleen, tonsils, and lymph nodes using in situ hybridization; within these tissues the mRNA was detected in lymphocytes, epithelial cells and vascular cells [812]. PRL receptors have been found in thymus, bone marrow, spleen, lymph nodes and peripheral macrophages and B lymphocytes [813]. A variety of cytokines affect pituitary PRL secretion and may be mediators, in part, of the PRL response to stress. In various experimental paradigms, interleukin-2 (IL-2) and IL-6 have been found to increase PRL secretion, and IL-1 and interferon-gamma inhibit PRL secretion [814–816]. However, in vivo these substances have little effect on PRL secretion in rats [816]. Conversely, PRL acts synergistically with IL-2 to induce maturation of nature killer cells [813]. Prolactin may have some clinically relevant effects on the immune system as well. Hyperprolactinemia caused by injections of ovine PRL into mice caused an increase in phagocytosis, intracellular killing of salmonella typhimurium, chemotaxis and overall survival [817]. Moreover, hyperprolactinemia caused by implanting pituitaries under the renal capsule caused increased autoimmune phenomena in the b/w mouse model [818]. In human patients with prolactinomas, polymorphonuclear cells show reduced chemotaxis [819,820] but monocytes display significantly increased bactericidal activity against Mycobacterium avium [821]. Yet other studies have shown that hyperprolactinemic patients display decreased natural killer T cell function and this effect is reversed with bromocriptine treatment [822]. Data on RPL levels in patients with various autoimmune disorders are conflicting. Nagy et al. [823] found a low PRL bioactivity to immunoreactivity ratio in patients with rheumatoid arthritis due to serum factors capable of inhibiting PRL bioactivity. In other studies hyperprolactinemia has been found in substantial portions of men and women with systemic lupus erythematosus [824–831]; in some of these studies PRL levels correlated with the extent of clinical disease and antinuclear antibody activity [826,828]. In many of these studies, no information is given regarding renal function, CNS involvement or medication use, so that whether the hyperprolactinemia is truly related to autoimmune disease or other factors known to elevate PRL levels is not clear. However, in several studies [827,828,831], lupus nephritis, hypothyroidism, medications and other known causes of hyperprolactinemia were specifically excluded. In

Chapter 4

one study of lupus in children, three of 33 had elevated PRL levels but all three had evidence of CNS lupus [829]. In an interesting report, Funauchi et al. [832] describe a patient with SLE and a prolactinoma whose serum PRL levels were significantly correlated to the serum anti-DNA titers and inversely correlated to the serum complement activity and whose flares and remission clinically correlated with stopping and resuming bromocriptine treatment. In studies of another inflammatory condition, Jara et al. [833] found PRL levels to be elevated in 36% of patients with Reiter’s syndrome. Furthermore, Bravo et al. found that four patients with Reiter’s syndrome had dramatic clinical improvement when treated with bromocriptine [834]. Giasuddin et al. [835] also reported elevated levels in patients with psoriasis. The clinical relevance of lymphoblastoid PRL and changes in immune function in normal individuals and those with hyperprolactinemia is still unclear. Although there is strong evidence in rodents that PRL is important in this regard, the evidence in humans is still very preliminary and what does exist is often contradictory and suggests that these effects of PRL are relatively minor compared to those of other immune regulators [836]. Nonetheless, the intriguing finding of increased PRL levels in many patients with autoimmune rheumatologic diseases merits continued study. PATHOLOGIC STATES OF PROLACTIN SECRETION Population studies of normal individuals reveal that PRL levels are not “normally” distributed and better fit a lognormal distribution contaminated by a small number of abnormally high measurements. Several studies have suggested that “elevated” PRL levels may be found in 1% to 10% of a random sample of individuals without evidence of endocrine abnormality [837–839]. When the frequency of abnormality in such a randomly sampled population is so high, it makes it difficult to determine what indeed is the upper limit of normal for that population and a number of mathematical formulations have been tried (for more complete discussions of this problem, see [837] and [840]). As discussed previously, PRL is secreted episodically, resulting in varying blood PRL levels throughout the day. Therefore, a PRL level in a single sample that is in the upper part of the normal range should be confirmed with one or two repeated tests before considering it to be abnormal.

Hypoprolactinemia As noted above, female mice with the PRL receptor KO are infertile with a number of defects occurring, including fewer primary follicles, fewer eggs ovulated, fewer eggs fertilized, impaired progression to the blastocyst stage, poor implantation and decreased estradiol and progesterone levels [679,688]. Male mice with the PRL receptor KO had only minimally impaired fertility [679] and those with the PRL gene KO had normal fertility [689].

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Idiopathic PRL deficiency has been described in only a single human [841]. This was a 30-year-old woman who had had a minimally delayed puberty (thelarche, age 15 and menarche, age 17), regular menses and normal body hair and breast development who conceived but was unable to lactate postpartum. Her basal PRL levels were all <3 ng/ml, during pregnancy her highest PRL level was 7.8 ng/ml and during the first few days postpartum her highest PRL level was 11.7 ng/ml. After TRH stimulation her PRL levels increased from 3.8 to 13.2 ng/ml. Her pituitary function was otherwise normal. PRL levels in her two children and other family members were normal. The ability of this woman to develop primary and secondary sexual characteristics normally and then conceive and deliver a child speak to the minimal, if any, importance of PRL in normal reproductive function, save that of postpartum lactation. No cases have been described analogous to the PRL gene or receptor KO mice. PRL deficiency may occur in the setting of panhypopituitarism, generally as a result of pituitary infarction. When the cause of the hypopituitarism is hypothalamic or stalk dysfunction, PRL levels generally rise due to disinhibition of PRL secretion (see below). However, when the pituitary tissue is actually destroyed, as in Sheehan’s peripartum necrosis, PRL levels are usually low [842]. In this latter circumstance, the hypoprolactinemia is manifest as an inability to lactate postpartum along with other symptoms of hypopituitarism. A deficiency of PRL may also occur in association with deficiencies of GH and TSH due to mutations in the Pit-1 and Prop-1 transcription factors necessary for tissuespecific PRL gene activation [46,47,49–51] (see above). PRL levels are also low in the condition of hypoparathyroidism [843]. Presumably this in some way reflects the importance of calcium as an intracellular mediator of PRL secretion, but the specific details of this have not been elucidated.

Hyperprolactinemia The differential diagnosis of sustained hyperprolactinemia is broad (Table 4.4). This section discusses those causes of hyperprolactinemia other than prolactinomas, which are discussed in Chapter 12. Medications

Neuroleptics The neuroleptic agents (phenothiazines and butyrophenones) uniformly result in elevated RPL levels, generally to less than 100 ng/ml but some patients have been reported with levels as high as 365 ng/ml [844,845]. Levels begin to rise within a few hours of starting therapy, continue to rise for 6 to 9 days, and then fall to lower, though still elevated levels after long-term treatment [844,845]. Chlorpromazine, sulpiride, and metoclopramide, major tranquilizers and antiemetics with potent dopamine antagonist action, have in fact been used as diagnostic agents to evaluate PRL secretory reserve. Neuroleptic agents raise PRL levels by blocking dopamine receptors, both on the

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Table 4.4.

Hypothalamic–Pituitary Function

Etiologies of hyperprolactinemia

Pituitary disease

Medications

Prolactinomas Acromegaly “Empty sella syndrome” Lymphocytic hypophysitis Cushing’s disease

Phenothiazines Haloperidol Monoamine-oxidase inhibitors Tricyclic antidepressants Reserpine Methyldopa Metoclopramide Amoxepin Cocaine Verapamil Serotonin reuptake inhibitors

Hypothalamic disease Craniopharyngiomas Meningiomas Dysgerminomas Nonsecreting pituitary adenomas Other tumors Sarcoidosis Histiocytosis X Neuraxis irradiation Vascular Pituitary stalk section Neurogenic Chest wall lesions Spinal cord lesions Breast stimulation

Other Pregnancy Hypothyroidism Chronic renal failure Cirrhosis Adrenal insufficiency Pseudocyesis Idiopathic

pituitary lactotrophs and in the hypothalamus. PRL levels usually fall to normal within 48 to 96 hours of discontinuation of neuroleptic drug therapy [844]. Combined serotonin/dopamine receptor antagonists such as risperidol cause similar elevations of PRL [846]. Antidepressants Tricyclic antidepressants cause modest hyperprolactinemia in about 25% of patients [847]. Monoamine oxidase (MAO) inhibitors, when used chronically, may also cause a minimal elevation of PRL levels [638]. The mechanisms by which these drugs cause increased PRL levels are not certain; they probably facilitate several possible stimulatory pathways and their effects on portal vessel dopamine are uncertain. Serotonin reuptake inhibitors, by increasing synaptic serotonin levels, commonly cause a mild increase in PRL to levels still within the normal range; rarely does hyperprolactinemia occur [489,848,849]. Opiates and Cocaine Chronic opiate abuse is associated with mild hyperprolactinemia and menstrual dysfunction (see above). Cocaine abuse has also been associated with chronic, mild hyperprolactinemia [850]. Antihypertensive Drugs Alpha-methyldopa causes moderate hyperprolactinemia [639] by reducing the effect level of dopamine that reaches the pituitary by inhibiting the enzyme l-aromatic amino acid decarboxylase, which is responsible for converting l-dopa to dopamine and by acting as a false neurotransmitter to decrease dopamine synthesis. Reserpine, a little-used antihypertensive drug now,

causes hyperprolactinemia by interfering with the storage of hypothalamic catecholamines in secretory granules [851]. ACE inhibitors, such as enalapril, facilitate PRL release in some individuals [582a] but sustained hyperprolactinemia causing symptoms has not been reported with their use. Calcium Channel Blockers Verapamil acutely and chronically, in most but not all experiments, has been found to increase basal PRL secretion and the PRL response to TRH [172,852,853] and patients have been described with galactorrhea associated with sustained hyperprolactinemia due to verapamil [171] (Fig. 4.15). In a survey of patients taking verapamil in an outpatient clinic, PRL levels were found to be elevated in 8.5% of patients [854]. Verapamil does not block the PRL inhibition caused by graded lowdose dopamine infusion, thereby showing lack of effect at the lactotroph [173]. However, it does block the hypothalamic generation of dopamine, as judged by finding decreased inhibition of PRL secretion by l-dopa after pretreatment with the peripheral dopa-decarboxylase inhibitor carbidopa, a maneuver that allows the assessment of central dopamine generation [173]. Interestingly, other calcium channel blockers such as the dihydropyridines and benzothiazepines have no action on PRL secretion, implying that the action of the phenylakylamine, verapamil, likely is acting on the neuronal N-type calcium channel [173]. Protease Inhibitors A recent report has shown that some HIV positive patients treated with protease inhibitors have developed galactorrhea and hyperprolactinemia [854a]. The mechanism for this is unknown at present. Stress

As noted previously, physical stress such as physical discomfort, exercise and hypoglycemia causes an acute, transient rise in PRL levels (see above). Chronic hyperprolactinemia due to prolonged physical stress has not been reported. Psychologic stress may cause minimal elevations of PRL [855] but chronic hyperprolactinemia has not been reported with any chronic psychiatric state except that of pseudocyesis, in which PRL levels fall with psychotherapy [856]. Renal disease

Hyperprolactinemia occurs in 73% to 91% of women and 25% to 57% of men with end-stage renal disease [857–859] (Fig. 4.16). PRL suppression by dopamine infusion is decreased [857] but this may be, at least in part, the result of delayed degradation of PRL in renal failure, since longterm use of bromocriptine results in suppression of PRL levels [858]. Although metabolic breakdown of PRL is delayed in renal failure, there is also increased production [264]. Therefore, there is disordered regulation of PRL secretion and possibly short-loop feedback, although the exact nature of the defect has not been defined. About onequarter of individuals with renal insufficiency not requiring dialysis (serum creatinine 2.0–12.0 ng/ml) have PRL levels in the 25–100 ng/ml range [800] (Fig. 4.17). When such

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FIGURE 4.15. (a) Effect of verapamil on the prolactin (PRL) response to a submaximal dose (100 mg) of thyrotropinreleasing hormone given at time 0. The open boxes represent the values before verapamil and the filled boxes represent the values after 1 week of verapamil (240 mg daily, sustained release). Note the elevated basal values. (b) Effect of verapamil on the suppression of the PRL by L-dopa (500 mg). The open boxes represent the values before verapamil and the filled boxes represent the values after 1 week of verapamil (240 mg daily, sustained release). Despite the elevated basal values after verapamil, the precentage suppressions were similar. From Kamal and Molitch [172]

FIGURE 4.16. Prolactin levels in patients requiring dialysis. CAPD, continuous ambulatory peritoneal dialysis. From Hou et al. [859]

FIGURE 4.17. Prolactin levels in patients with renal insufficiency, including patients on medications (M, metoclopramide; all others designated receiving medications were taking methyldopa). From Hou et al. [859]

has occurred in some women treated with bromocriptine [858], but reports of restored ovulation are rare [857]. patients take medications known to alter hypothalamic regulation of PRL, such as methyldopa or metoclopramide, PRL levels may rise to over 2000 ng/ml [859]. Correction of the renal failure with transplantation causes a return of PRL levels to normal [857,859]. Hyperprolactinemia plays a role in the hypogonadism of chronic renal failure, but probably does not explain all of the abnormalities in every case. Return of normal menses

Cirrhosis

Basal PRL levels are increased in patients with alcoholic cirrhosis in frequencies varying from 16% to 100% [860–862] and in patients with nonalcoholic cirrhosis from 5% [863] to 13% [860]. In a study by Van Thiel et al. [861] the PRL response to TRH was normal in 75% of the hyperprolactinemic men with alcoholic cirrhosis and blunted in

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the remainder. In another study, PRL responses to TRH were greatly augmented in patients with a variety of types of cirrhosis [864]. In a third study, 50% of patients with hepatic encephalopathy were found to be hyperprolactinemic [865]. Although l-dopa caused a decrease in PRL levels in all, inhibition of peripheral decarboxylase with carbidopa resulted in an impaired PRL response to l-dopa, implying an underlying defect in hypothalamic dopamine generation as the cause of the hyperprolactinemia in these encephalopathic patients. In women without cirrhosis, acute alcohol withdrawal is associated with hyperprolactinemia in 50% [866]. Hypothyroidism

Primary hypothyroidism is associated with a modest increase in the level of PRL in 40% of patients, but levels greater than 25 ng/ml are reached in only 10% [867]. The mechanisms involved probably include increased TRH production, increased sensitivity of lactotrophs to TRH, and possibly increased pituitary VIP generation (see above). Because many patients with long-standing hypothyroidism may have evidence of pituitary enlargement on x-ray, the finding of hyperprolactinemia, galactorrhea and/or amenorrhea associated with an enlarged pituitary seen in hypothyroidism may be easily confused with a prolactinoma. Therapy with Lthyroxine will cause the PRL levels to return to normal and can even result in a regression of pituitary size [868]. Adrenal Insufficiency

As discussed above, glucocorticoids have a suppressive effect on PRL gene transcription and PRL release. Adrenalectomy has been reported to cause hyperprolactinemia in animals [869]. Rare cases in humans have been reported of hyperprolactinemia occurring in patients with adrenal insufficiency in whom the PRL levels return to normal with glucocorticoid replacement [870–874]. Neurogenic

As discussed above, sexual breast stimulation and suckling cause a reflex release of PRL that is mediated, in part, by afferent neural pathways going through the spinal cord. Chest wall and cervical cord lesions have been reported to result in elevated PRL levels and galactorrhea through stimulation of these afferent neural pathways [875,876]. Similar chronic elevations of PRL have been reported after mastectomy and thoracotomy [877] and with chronic spinal cord injuries [878]. Ectopic Prolactin Secretion

Ectopic production of PRL is exceedingly rare. In a careful evaluation of 215 patients with a variety of malignancies, PRL elevations were found in only 15 and in 12 of these the elevations could be explained by the use of phenothiazines, opiates, or prior irradiation to the chest wall or head [879]. In one of the remaining three patients subsequent samples all had normal PRL measurements and in the other two, the cancer (lung and breast) could also have caused the

modest hyperprolactinemia by stimulation of chest wall afferent nerves. However in another series of patients with uterine cervical carcinoma, 229 out of 743 patients had elevated serum PRL levels; surgery in 86 of these patients resulted in normalization of PRL levels [880]. In this same study, 22 out of 49 cervical carcinomas stained positively for PRL by immunohistochemistry and 5 out of 8 carcinomas produced PRL when cultured [880]. In none of these cases were all of these aspects put together, i.e. elevated serum levels that normalized after surgery and whose tumors both stained for PRL and produced PRL in vitro. However, symptomatic hyperprolactinemia due to well-documented PRL production from a renal cell carcinoma [881], a gonadoblastoma [882] and ectopic pituitary tissue in two ovarian teratomas [883,884] have been reported. Given the great frequency of prolactinomas, “idiopathic hyperprolactinemia” and other causes of hyperprolactinemia, a search for an ectopic source of PRL secretion is not warranted unless some other tumor shows up coincidentally. Hypothalamic/Pituitary Stalk Disease

Hyperprolactinemia caused by lesions of the hypothalamus and of the pituitary stalk is due to disturbance of the neuroendocrine mechanisms that control PRL secretion. From hypothalamic lesion work in animals, it has been assumed that this PRL elevation is due to disinhibition of the tonic PIF (dopamine) acting at the level of the pituitary lactotrophs. PRL secretory dynamics were studied systematically in a group of such patients with hypothalamic hyperprolactinemia (six with hypothalamic lesions such as craniopharyngiomas, eosinophilic granuloma), eight with large “nonfunctioning” pituitary adenomas with marked suprasellar extension, and seven with partially empty sellas [885]. Basal PRL levels were <100 ng/ml in all but 1 (Fig. 4.18). However, in one patient with a Rathke’s cleft cyst, preoperative PRL levels ranged between 93 and 148 ng/ml; she had normal TSH and ACTH function but following transsphenoidal resection she developed diabetes insipidus and panhypopituitarism, indicating high stalk section, and PRL levels fell to 25–28 ng/ml. Interestingly, of seven of these patients tested with insulin-induced hypoglycemia, three had normal PRL responses and seven out of 12 showed a significant inhibition in response to L-dopa even after peripheral conversion to dopamine was blocked by carbidopa pretreatment. Furthermore, most of these patients had normal TSH and ACTH function. These results imply that there still is significant transmission of hypothalamic releasing factors to the pituitary in most of these cases, despite increased PRL levels. The fall in PRL levels in the patient with the Rathke’s cleft cyst, along with the development of a true stalk section syndrome, implies that there is a difference in patients with total stalk section in whom the PRL rise is due solely to dopamine deficiency and those with partial stalk/hypothalamic dysfunction who have dopamine deficiency plus continued PRF activity, resulting in higher PRL levels. In addition to this case with a Rathke’s

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FIGURE 4.18. Basal prolactin levels in patients with hypothalamic disease. From Molitch and Reichlin [900]

cleft cyst with a basal PRL level of 122 ng/ml a number of cases have been reported in the literature by others with PRL levels between 104 and 219 ng/ml [886–891]. A case of a patient with a PRL of 662 ng/ml whose tumor did not stain for PRL has been well-documented [892], but cases with such high PRL levels must be extraordinarily rare. It is also possible that some cases of hyperprolactinemia due to hypothalamic disease could be due to increased PRF activity with normal PIF activity. Anterior hypothalamic lesions that are irritative could potentially stimulate PRF pathways, similar to findings in experimental animals [893,894]. The findings of a normal suppression of PRL with l-dopa alone and combined with carbidopa in two of our patients supports the concept of an intact TIDA system in these two patients [885]. Idiopathic Hyperprolactinemia

When no specific cause is found with the evaluation outlined below, the hyperprolactinemia is of uncertain etiology and has been designated to be idiopathic. It is recognized that in many such cases small prolactinomas that are too small to be detected by current radiologic techniques may be present. In other cases the hyperprolactinemia is due to presumed hypothalamic regulatory dysfunction, but no dysfunction specific to idiopathic hyperprolactinemia has been definitively elucidated. Long-term follow-up of such patients has found that in about one-third, PRL levels return to normal, in 10% to 15% there is a rise in PRL levels to >50% over baseline, and in the remaining patients prolactin levels remain stable [895,896]. Over a 2 to 6 year followup of 199 patients, only 23 developed evidence of microadenomas and none developed macroadenomas [895–899]. CLINICAL TESTING As mentioned above, PRL is secreted episodically and some PRL levels during the day may be above the upper limit of

normal established for a given laboratory. Thus the finding of minimally elevated levels in blood requires confirmation in several samples. As indicated in the previous section, there are a number of conditions that may cause moderate PRL elevations, although generally to levels <250 ng/ml. A careful history and physical examination, screening blood chemistries, thyroid function tests and a pregnancy test will exclude virtually all causes except for hypothalamic– pituitary disease. When there is no obvious cause of the hyperprolactinemia from the routine screening, a radiologic evaluation of the hypothalamic–pituitary area is mandatory to exclude a mass lesion. This includes patients with even mild PRL elevations. At present, such an evaluation is generally done using magnetic resonance imaging (MRI) with gadolinium enhancement or computed tomography (CT) with intravenous contrast enhancement (see Chapter 20). It should be emphasized here that it is very important to distinguish between a large nonsecreting tumor causing modest PRL elevations (usually <250 ng/ml, see above) from a PRL-secreting macroadenoma (PRL levels usually >>250 ng/ml), as the therapy may be quite different. PRLsecreting macroadenomas generally will respond quite readily to bromocriptine and other dopamine agonists with size reduction (see Chapter 11, p. 425) whereas only about 10% of nonsecreting pituitary tumors respond in this manner. Stimulation and suppression tests give nonspecific results. Although blunted PRL responses to TRH (Fig. 4.19) and hypoglycemia (Fig. 4.20) are characteristic of patients with prolactinomas, such blunted responses are also seen in patients with hypothalamic dysfunction and other causes of hyperprolactinemia and normal PRL responses may be seen in 10% to 15% of patients with prolactinomas [900–902]. Similar nonspecific results have been found with chlorpromazine, metoclopramide, nomifensine, and l-dopa and also carbidopa [900–902]. Thus, consensus has developed that such stimulation and suppression tests are worthless in the

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FIGURE 4.19. Prolactin responses to thyrotropin-releasing hormone in patients with hyperprolactinemia of different etiologies. Note the marked blunting of the responses of all categories of hyperprolactinemic patients in comparison to those of normal controls. From Molitch and Reichlin [900]

FIGURE 4.20. Prolactin responses to insulin-induced hypoglycemia in patients with hyperprolactinemia of different etiologies. Note the marked blunting of the responses of all categories of hyperprolactinemic patients in comparison to those of normal controls. From Molitch and Reichlin [900]

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168 Di Renzo G, Amoroso S, Maida P et al. Effect of different Ca2+ entry blockers on dopamine-induced inhibition of in vitro prolactin secretion. Eur J Pharmacol 1988;146:201–206. 169 Kuan SI, Login IS, Judd AM. A comparison of the concentration-dependent actions of thyrotropin-releasing hormone, angiotensin II, bradykinin. Endocrinology 1990;127:1841–1848. 170 Schrey MP, Clark HJ, Franks S. The dopaminergic regulation of anterior pituitary 45Ca2+ homeostasis and prolactin secretion. Endocrinology 1986;108: 423–429. 171 Gluskin LE, Strasberg B, Shah JH. Verapamil-induced hyperprolactinemia and galactorrhea. Ann of Intern Med 1981;95:66–67. 172 Kamal TJ, Molitch ME. Effects of calcium channel blockade with verapamil on the prolactin responses to TRH, L-DOPA and bromocriptine. Am J Med Sci 1992;304:289–293. 173 Kelley SR, Kamal TJ, Molitch ME. Mechanism of verapamil calcium channel blockade-induced hyperprolactinemia. Am J Physiol 1996;270:E96– E100. 174 Sinha YN, Gillian TA, Lee DW et al. Cleaved prolactin: evidence for its occurrence in human pituitary gland and plasma. J Clin Endocrinol Metab 1985;60:239–243. 175 Mittra I. A novel “cleaved prolactin” in the rat pituitary. I. Biosynthesis, characterization and regulatory control. Biochem Biophys Res Commun 1980;95:1750–1759. 176 Sinha YN, Gilligan TA. A cleaved form of prolactin in the mouse pituitary gland: identification and comparison of in vitro synthesis and release in strains with high and low incidences of mammary tumors. Endocrinology 1984;114: 2046–2050. 177 Mittra I. A novel “cleaved prolactin” in the rat pituitary. II. In vivo mammary mitogenic activity of its N-terminal 16K moiety. Biochem Biophys Res Commun 1980;95:1760–1767. 178 Oetting WS, Walker AM. Intracellular processing of prolactin. Endocrinology 1985;117:1565–1570. 179 Sinha YN, Jacobsen BP. Structural and immunologic evidence for a small molecular weight (“21K”) variant of prolactin. Endocrinology 1988;123: 1364–1370. 180 Oetting WS, Ho TWC, Greenan JR, Walker AM. Production and secretion of the 21–23.5 kDa prolactin-like molecules. Mol Cell Endocrinol 1989;61: 189–199. 181 Greenan JR, Balden E, Ho TWC, Walker AM. Biosynthesis of the secreted 24 K isoforms of prolactin. Endocrinology 1989;125:2041–2047. 182 Suh HK, Frantz AG. Size heterogeneity of human prolactin in plasma and pituitary extracts. J Clin Endocrinol Metab 1974;39:928–935. 183 Guyda HJ. Heterogeneity of human growth hormone and prolactin secreted in vitro: immunoassay and radioreceptor assay correlations. J Clin Endocrinol Metab 1975;41:953–967. 184 Garnier PE, Aubert ML, Kaplan ASL, Grumbach MM. Heterogeneity of pituitary and plasma prolactin in man: Decreased affinity of “big” prolactin in a radioreceptor assay and evidence for its secretion. Endocrinology 1978;47: 1273–1279. 185 Wallis M, Daniels M, Ellis SA. Size heterogeneity of rat pituitary prolactin. Biochem J 1980;189:605–614. 186 Whitaker MD, Klee GG, Kao PC et al. Demonstration of biological activity of prolactin molecular weight variants in human sera. J Clin Endocrinol Metab 1983;58:826–830. 187 Torres AI, Aoki A. Release of big and small molecular forms of prolactin: dependence upon dynamic state of the lactotroph. J Endocrinology 1987;114: 213–220. 188 Lawson DM, Gala RR, Chin ML, Haisenleder DH. Size heterogeneity of plasma prolactin in the rat: TRH and serotonin-induced changes. Life Sci 1980;27:1147–1151. 189 Molitch ME, Raiti S, Baumann G et al. Pharmacokinetic studies of highly purified human prolactin normal human subjects. J Clin Endocrinol Metab 1987;65:299–304. 190 Andino NA, Bidot C, Valdez M, Machado AJ. Chromatographic pattern of circulating prolactin in ovulatory hyperprolactinemia. Fertil Steril 1985;44: 600–605. 191 Larrea F, Villanueva C, Cravioto MC. Further evidence that Big, Big prolactin is preferentially secreted in women with hyperprolactinemia and normal ovarian function. Fertil Steril 1985;44:25–30. 192 Jackson RD, Wortsman J, Malarkey WB. Characterization of a large molecular weight prolactin in women with idiopathic hyperprolactinemia and normal menses. J Clin Endocrinol Metab 1985;61:258–264. 193 Wortsman J, Carlson HE, Malarkey WB. Macroprolactinemia as the cause of elevated serum prolactin in men. Am J Med 1989;86:704–706.

194 Corenblum B. Asymptomatic hyperprolactinemia resulting from macroprolactinemia. Fertil Steril 1990;53(1):165–167. 195 Vieira JGH, Tachibana TT, Obara LH, Maciel RMB. Extensive experience and validation of polyethylene glyco precipitation as a screening method for macroprolactinemia. Clin Chem 198;44:758–759. 196 Lewis UJ, Singh RNP, Sinha YN, Vanderlaan WP. Glycosylated human prolactin. Endocrinology 1985;116:359–363. 197 Lewis UJ, Singh RNP, Lewis LJ. Two forms of glycosylated human prolactin have different pigeon crop sac-stimulating activities. Endocrinology 1989;24: 1558–1563. 198 Markoff E, Lee DW. Glycosylated prolactin is a major circulating variant in human serum. Endocrinology 1987;65:1102–1106. 199 Hashim IA, Aston R, Butler J. The proportion of glycosylated prolactin in serum is decreased in hyperprolactinemic states. J Clin Endocrinol Metab 1990;71:111–115. 200 Haro LS, Lee DW, Singh RNP et al. Glycosylated human prolactin: alterations in glycosylation pattern modify affinity for lactogen receptor and values in prolactin radiommunoassay. J Clin Endocrinol Metab 1990;71:379–383. 201 Markoff E, Sigel MB, Lacour N et al. Glycosylation selectively alters the biological activity of prolactin. Endocrinology 1988;123:1303–1306. 202 Pellegrini I, Gunz G, Ronin C et al. Polymorphism of prolactin secreted by human prolactinoma cells: immunological, receptor binding and biological properties of the glycosylated and nonglycosylated forms. Endocrinology 1988;122:2667–2674. 203 Young KH, Buhi WC, Horseman N et al. Biological activities of glycosylated and nonglycosylated porcine prolactin. Mol Cell Endocrinol 1990;71:155–162. 204 Oetting WS, Tuazon PT, Traugh JA, Walker AM. Phosphorylation of prolactin. J Biol Chem 1986;261:1649–1652. 205 Brooks CL, Kim BG, Aphale P et al. Phosphorylated variant of bovine prolactin. Mol Cell Endocrinol 1990;71:117–123. 206 Soares MJ, Müller H, Orwig KE et al. The uteroplacental prolactin family and pregnancy. Biol Reprod 1998;58:273–284. 207 Linzer DIH, Fisher SJ. The placenta and the prolactin family of hormones: regulation of the physiology of pregnancy. Mol Endocrinol 1999;13:837–840. 208 Tyson JE, Friesen HG. Factors influencing the secretion of human prolactin and growth hormone in menstrual and gestational women. Am J Obstet Gynecol 1973;116:377–387. 209 Rigg LA, Lein A, Yen SSC. Pattern of increase in circulating prolactin levels during human gestation. Am J Obstet Gynecol 1977;129:454–456. 210 Ben-David M, Rodbard D, Bates RW, Bridson E, Chrambach A. Human prolactin in plasma, amniotic fluid and pituitary: identity and characterization by criteria of electrophoresis and isoelectric focusing in polyacrylamide gel. J Clin Endocrinol Metab 1973;36:951–964. 211 Rathman P, Cederqvist L, Saxena BB. Isolation of prolactin from human amniotic fluid. Biochem Biophys Acta 1977;492:186–193. 212 Friesen H, Hwang P, Guyda H et al. In: Boyns AR, Griffiths K, eds. Prolactin and Carcinogenesis. Wales: Alpha Omega Publishing, 1972:64–80. 213 Riddich DH, Kusmik WF. Decidua: a possible source of amniotic fluid prolactin. Am J Obstet Gynecol 1977;127:187–192. 214 Golander A, Hurley T, Barrett J et al. Prolactin synthesis by human choriondecidual tissue: A possible source of prolactin in the amniotic fluid. Science 1978;202:311–313. 215 De Ziegler D, Gurpide E. Production of prolactin by cultures of cells from human decidua. J Clin Endocrinol Metab 1982;55:511–515. 216 Tomita K, McCoshen JA, Friesen HG, Tyson JE. Quantitative comparison between biological and immunological activities of prolactin derived from human fetal and maternal sources. J Clin Endocrinol Metab 1982;55:269–271. 217 Takahashi H, Nabeshima Y, Nabeshima Y-I et al. Molecular cloning and nucleotide sequence of DNA complementary to human decidual prolactin mRNA. J Biol Chem 1984;95:1491–1499. 218 Huang JR, Tseng L, Bischof P, Janne OA. Regulation of prolactin production by progestin, estrogen, and relaxin in human endometrial stromal cells. Endocrinology 1987;121:2011–2017. 219 Gellersen B, DiMattia GE, Friesen HG, Bohnet HG. Prolactin (PRL) mRNA from human decidua differs from pituitary PRL mRNA but resembles the IM-9-P3 lymphoblast PRL transcript. Mol Cell Endocrinol 1989;64:127–130. 220 Lee DW, Markoff E. Synthesis and release of glycosylated prolactin by human decidua in vitro. J Clin Endocrinol Metab 1986;62:990–994. 221 Heffner LJ, Gramates LS, Yuan RW. A glycosylated prolactin species is covalently bound to immunoglobulin in human amniotic fluid. Biochem Biophys Res Comm 1989;165:299–305. 222 Jayatilak PG, Glaser LA, Busuray R et al. Identification and partial characterization of a prolactin-like hormone produced by rat decidual tissue. Proc Natl Acad Sci USA 1985;82:217–221.

Chapter 4 223 Herz Z, Khan I, Jayatilak PG, Gibori G. Evidence for the secretion of decidual luteotropin: a prolactin-like hormone produced by rat decidual cells. Endocrinology 1986;118:2203–2209. 224 Jayatilak PG, Puryear TK, Herz Z et al. Protein secretion by mesometrial and antimesometrial rat decidual tissue: Evidence for differential gene expression. Endocrinology 1989;125:659–666. 225 Bigazzi M, Ronga R, Lancranjan I et al. A pregnancy in an acromegalic woman during bromocriptine treatment: effects on growth hormone and prolactin in the maternal, fetal and amniotic compartments. J Clin Endocrinol Metab 1979;48:9–12. 226 Kubota T, Nagae M, Yoshimasa Y et al. Prolactin-releasing system in maternal, fetal and amniotic compartments during labor. Obstetrics and Gynecology 1986;68:80–86. 227 Maslar IA, Ansbacher R. Effect of short-duration progesterone treatment on decidual prolactin by cultures of proliferative human endometrium. Fertil Steril 1988;50:250–254. 228 Golander A, Richards R, Thrailkill K et al. Decidual prolactin (PRL) releasing factors stimulate the synthesis of PRL from human decidual cells. Endocrinology 1988;123:335–340. 229 Thrailkill KM, Golander A, Underwood LE, Handwerger S. Insulin-like growth factor I stimulates the synthesis and release of prolactin from human decidual cells. Endocrinology 1988;123:2930–2934. 230 Thrailkill KM, Golander A, Underwood LE et al. Insulin stimulates the synthesis and release of prolactin from human decidual cells. Endocrinology 1989;124:3010–3014. 231 Josimovich JB, Merisko K, Boccella L. Amniotic prolactin control over amniotic and fetal extracellular fluid water and electrolytes in the rhesus monkey. Endocrinology 1977;100:564–570. 232 Johnson JWC, Tyson JE, Mitzner W et al. Amniotic fluid prolactin and fetal lung maturation. Am J Obstet Gynecol 1985;153:372–375. 233 Horrobin DF. Prolactin as a regulator of fluid and electrolyte metabolism in mammals. Fed Proc 1980;39:2567–2570. 234 Di Mattia GE, Gellersen B, Duckworth ML. Human prolactin gene expression. J Biol Chem 1990;265:16412–16421. 235 Di Mattia GE, Gellersen B, Bohnet HG, Friesen HG. A human Blymphoblastoid cell line produces prolactin. Endocrinology 1988;122:2508–2517. 236 Gellersen B, Di Mattia GE, Friesen HG. Regulation of prolactin secretion in the human B-lymphoblastoid cell line IM-9-P3 by dexamethasone but not other regulators of pituitary prolactin secretion. Endocrinology 1989;125: 2853–2861. 237 Pelletier G, Lemay A, Beraud G, Labrie F. Ultrastructural changes accompanying the stimulatory effect of n-monobutyryl adenosine 3¢,5¢monophosphate on the release of growth hormone (GH), prolactin (PRL) and adrenocorticotropic hormone (ACTH) in rat anterior pituitary gland in vitro. Endocrinology 1972;91:1355–1371. 238 Walker AM, Farquhar MG. Preferential release of newly synthesized prolactin granules is the result of functional heterogeneity among mammotrophs. Endocrinology 1980;107:1095–1104. 239 Nicoll CS, Swearingen KC. Preliminary observations on prolactin and growth hormone turnover in rat adenohyphyses in vivo. In: Martin L, Motta M, Fraschini F, eds. The Hypothalamus. New York: Academic Press, 1970; 449–462. 240 Swearingen KC, Nicoll CS. Prolactin turnover in rat adenohypophyses in vivo: Its evaluation as a method for estimating secretion rates. Endocrinology 1972;53:1–15. 241 Swearingen KC. Heterogeneous turnover of adenohypophysial prolactin. Endocrinology 1971;89:1380–1388. 242 Stachura ME. Sequestration of an early-release pool of growth hormone and prolactin in GH3 rat pituitary tumor cells. Endocrinology 1982;111:1769– 1777. 243 Stachura ME, Tyler JM, Kent PG. Pituitary immediate release pools of growth hormone and prolactin are preferentially refilled by new rather stored hormone. Endocrinology 1989;125:444–449. 244 Jacobs LS, Lorenson MY. Cysteamine, zinc and thiols modify detectability of rat pituitary prolactin: a comparison with effects on bovine prolactin suggests differences in hormone storage. Metabolism 1986;35:209–215. 245 Sagar SM, Millard WJ, Martin JB, Murchison SC. The mechanism of action of cysteamine in depleting prolactin immunoreactivity. Endocrinology 1985; 117:591–600. 246 Lorenson MY, Robson DL, Jacobs LS. Detectability of pituitary PRL and GH by immunoassay is increased by thiols and suppressed by divalent cations. Endocrinology 1983;112:1880–1882. 247 Scammell JG, Dannies PS. Depletion of pituitary prolactin by cysteamine is due to loss of immunological activity. Endocrinology 1984;114:712–716.

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248 Mena F, Clapp C, Aguayo D et al. Thiol regulation of depletiontransformation and release of prolactin by the pituitary of the lactating rat. Endocrinology 1986;118:1795–1802. 249 Copeland PM, Martin JB, Ridgway EC. Cysteamine decreases prolactin responsiveness to thyrotropin-releasing hormone in normal men. Am J Med 1986;291:16–19. 250 Brown M, Fisher L, Mason RT et al. Neurobiological actions of cysteamine. Fed Proc 1985;44:2556–2560. 251 Das Gaines RE, Cotes PM. International reference preparation of human prolactin for immunoassay: definition of the international unit, report of a collaborative study and comparison of estimates of human prolactin made in various laboratories. J Endocrinol 1979;80:157–168. 252 Schulster D, Das Gaines RE, Jeffcoate SL. International standards for human prolactin: Calibration by international collaborative study. J Endocrinol 1989;121:157–166. 253 Fujimoto VY, Clifton DK, Cohen NL, Soules MR. Variability of serum prolactin and progesterone levels in normal women: The relevance of single hormone measurements in the clinical setting. Obst and Gynecol 1990;76: 71–78. 254 St-Jean E, Blain F, Comtois R. High prolactin levels may be missed by immunoradiometric assay in patients with macroprolactinomas. Clin Endocrinol 1996;44:305–309. 255 Barkan A, Chandler WF. Giant pituitary prolactinoma with falsely low serum prolactin: the pitfall of the “high-dose hook effect”: case report. Neurosurgery 1998;9:13–15. 256 Kleinberg DL, Frantz AG. Human prolactin: Measurement in plasma by in vitro bioassay. J Clin Invest 1971;50:1557–1568. 257 Turkington RW. Measurement of prolactin activity in human serum by the induction of specific milk proteins in mammary gland in vitro. J Clin Endocrinol Metab 1971;33:210–217. 258 Shiu RPC, Kelly PA, Friesen HG. Radioreceptor assay for prolactin and other lactogenic hormones. Science 1973;180:968–970. 259 Tanaka T, Shiu RPC, Gout PW et al. A new sensitive and specific bioassay for lactogenic hormones: measurement of prolactin and growth hormone in human serum. J Clin Endocrinol Metab 1980;51:1058–1063. 260 Tanaka T, Shishiba Y, Gout PW, Beer CT. Radioimmunoassay and bioassay of human growth hormone and Human prolactin. J Clin Endocrinol Metab 1983;56:18–20. 261 Rowe RC, Cowden EA, Faiman C, Friesen HG. Correlation of Nb2 bioassay and radioimmunoassay values for human serum prolactin. J Clin Endocrinol Metab 1983;57:942–946. 262 Frawley LS, Clark CL, Schoderbek WC. A novel bioassay for lactogenic activity. Demonstration that prolactin cells differ from one another in bio- and immuno-potencies of secreted hormone. Endocrinology 1986;119:2867–2869. 263 Cooper BS, Ridgway EC, Kliman B. Metabolic clearance and production rates of prolactin in man. J Clin Invest 1979;64:1669–1680. 264 Sievertsen GD, Lim VS, Nakawatase C, Frohman LA. Metabolic clearance and secretion rates of human prolactin in normal subjects and in patients with chronic renal failure. J Clin Endocrinol Metab 1980;50:846–852. 265 Kapcala LP, Molitch ME, Arno J et al. Twenty-four hour prolactin secretory patterns in women with galactorrhea, normal menses, normal random prolactin levels and abnormal sellar tomograms. J Endocrinol Invest 1984;7:455–460. 266 Bratusch-Marrain P, Bjorkman O. Hepatic disposal of endogenous growth hormone and prolactin in man. Eur J Clin Invest 1979;9:257–260. 267 Falconer IR, Vacek AT. Degradation of 125I-labeled prolactin in the rabbit: effect of nephrectomy and prolactin infusion. J Endocrinol 1983;99:369–377. 268 Shin SH, Reifel CW. Adenohypophysis has an inherent property for pulsatile prolactin secretion. Neuroendocrinology 1981;32:139–144. 269 Stewart JK, Clifton DK, Koerker DJ, Rogol AD. Pulsatile release of growth hormone and prolactin from the primate pituitary in vitro. Endocrinology 1985;116:1–5. 270 Veldhuis JD, Johnson, L. Operating characteristics of the hypothalamo–pituitary–gonadal axis in men: circadian, ultradian, and pulsatile release of prolactin and its temporal coupling with luteinizing hormone. J Clin Endocrinol Metab 1988;67:116–123. 271 Greenspan SL, Klibanski A, Rowe JW. Age alters pulsatile prolactin release: influence of dopaminergic inhibition. Am J Physiol 1990;258:E799–E804. 272 Samuels MH, Henry P, Kleinschmidt-DeMasters B et al. Pulsatile prolactin secretion in hyperprolactinemia due to presumed pituitary stalk interruption. J Clin Endocrinol Metab 1991;73:1289–1293. 273 Parker DC, Rossman LG, Vanderlaan EF. Relation of sleep-entrained human prolactin release to REM-NonREM cycles. J Clin Endocrinol Metab 1973;38: 646–651.

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274 Sassin JF, Frantz AG, Kapen S, Weitzman ED. The nocturnal rise of human prolactin is dependent upon sleep. J Clin Endocrinol Metab 1973;37:436– 440. 275 Stern JM, Reichlin S. Prolactin circadian rhythm persists throughout lactation in women. Neuroendocrinology 1990;51:31–37. 276 Linkowski P, Spiegel K, Kerkhofs M et al. Genetic and environmental influences on prolactin secretion during wake and during sleep. Am J Physiol 1998;274:E909–E919. 277 Weitzman ED, Boyar RM, Kapen S. The relationship of sleep and sleep stages of neuroendocrine secretion and biological rhythms in man. Rec Prog Horm Res 1975;31:399–446. 278 Quigley ME, Ropert JF. Acute prolactin release triggered by feeding. J Clin Endocrinol Metab 1981;52:1043–1045. 279 Carlson HE, Wasser HL, Levin SR. Prolactin stimulation by meals is related to protein content. J Clin Endocrinol Metab 1983;57:334–338. 280 Carlson HE. Prolactin stimulation by protein is mediated by amino acids in humans. J Clin Endocrinol Metab 1989;69:7–14. 281 Carlson HE, Miglietta JT, Roginsky MS. Stimulation of pituitary hormone secretion by neurotransmitter amino acids in humans. Metabolism 1989;38:1179–1182. 282 Carlson HE, Hyman DB, Blitzer MG. Evidence for an intracerebral action of phenylalanine in stimulation of prolactin secretion: Interaction of large neutral amino acids. J Clin Endocrinol Metab 1990;70:814–816. 283 Poindexter AN, Buttram VC, Besch P. Circulating prolactin levels, I. Normal females. Int J of Fertil 1977;22:1–5. 284 Vekemans M, Robyn C. Influence of age on serum prolactin levels in women and men. Brit Med J 1975;2:738–739. 285 Djursing H, Hagen C, Moller J. Short and long term fluctuations in plasma prolactin concentration in normal subjects. Acta Endocrinol 1981;97:1–6. 286 Rolandi E, Magnani G, Sannia A. Evaluation of PRL secretion in elderly subjects. Acta Endocrinol 1982;100:351–357. 287 Balint-Peric´ LA, Prelevic´ GM. Changes on prolactin levels with the menopause: the effects of estrogen/androgen and calcitonin treatment. Gynecol Endocrinol 1997;11:275–280. 288 Foth D, Römer T. Prolactin serum levels in postmenopausal women receiving long-term hormone replacement therapy. Gynecol Obstet Invest 1997;44: 124–126. 289 Touraine P, Deneux C, Plu-Bureau G et al. Hormonal replacement therapy in menopausal women with a history of hyperprolactinemia. J Endocrinol Invest 1998;21:732–736. 290 Iranmesh A, Mulligan T, Veldhuis JD. Mechanisms subserving the physiological nocturnal relative hypoprolactinemia of healthy older men: dual decline in prolactin secretory burst mass and basal release with preservation of pulse duration, frequency, and interpulse interval—a General Clinical Research Center Study. J Clin Endocrinol Metab 1999;84:1083–1090. 291 McNeilly AS, Chard T. Circulating levels of prolactin during the menstrual cycle. Clin Endocrinol 1974;3:105–112. 292 Franchimont P, Dourcy C, Legros JJ. Prolactin levels during the menstrual cycle. Clin Endocrinol 1976;5:643–650. 293 Cole EN, England PC, Sellwood RA. Serum prolactin concentrations throughout the menstrual cycle of normal women and patients with recent breast cancer. Eur J Cancer 1977;13:677–684. 294 Gordon JS, Wu CH, Mikhail G. Daily plasma prolactin in various gynecologic endocrinopathies. Fertil Steril 1979;31:385–391. 295 Braund W, Roeger DC, Judd SJ. Synchronous secretion of luteinizing hormone and prolactin in the human luteal phase: neuroendocrine mechanisms. J Clin Endocrinol Metab 1984;58:293–297. 296 Noel GL, Suh HK, Frantz AG. Prolactin release during nursing and breast stimulation in postpartum and nonpostpartum subjects. J Clin Endocrinol Metab 1974;38:413–423. 297 Lee LR, Haisenleder DJ, Marshall JC. The role of the suckling stimulus in regulating pituitary prolactin mRNA in the rat. Mol Cell Endocrinol 1989; 64:243–249. 298 Howie PW, McNeilly AS, McArdle T, Smart L. The relationship between suckling-induced prolactin response and lactogenesis. J Clin Endocrinol Metab 1980;50:670–673. 299 Weitzman RW, Leake RD, Rubin RT. The effect of nursing on neurohypophyseal hormone and prolactin secretion in human subjects. J Clin Endocrinol Metab 1980;51:836–839. 300 Johnston JM, Amico JA. A prospective longitudinal study of the release of oxytocin and prolactin in response to infant suckling in long term lactation. J Clin Endocrinol Metab 1986;62:653–657. 301 Bonnar J, Franklin M, Nott PN, McNeilly AS. Effect of breast-feeding on pituitary–ovarian function after childbirth. Brit Med J 1975;4:82–84.

302 Battin DA, Marrs RP, Fleiss PM. Effect of suckling on serum prolactin, luteinizing hormone, follicle-stimulating hormone, and estradiol during prolonged lactation. Obstet Gynecol 1985;65:785–788. 303 Stern JM, Konner M, Herman TN, Reichlin S. Nursing behavior, prolactin and postpartum amenorrhoea during prolonged lactation in American and !Kung mothers. Clin Endocrinol 1986;25:247–258. 304 Delvoye P, Demaegd M, Uwayitu-Nyampeta, Robyn C. Serum prolactin, gonadotropins, and estradiol in menstruating and amenorrheic mothers during two years’ lactation. Am J Obstet Gynecol 1978;130:635–639. 305 Prema K, Naidu AN, Kumari SN. Lactation and fertility. Am J Clin Nutr 1979;32:1298–1303. 306 Konner M, Worthman C. Nursing frequency, gonadal function, and birth spacing among !Kung hunter-gatherers. Science 1980;207:788–792. 307 Kolodny RC, Jacobs LS, Daughaday WH. Mammary stimulation causes prolactin secretion in non-lactating women. Nature 1972;238:284–286. 308 Neill JD. Effect of “stress” on serum prolactin and luteinizing hormone levels during the estrous cycle of the rat. Endocrinology 1970;87:1192–1197. 309 Noel GL, Suh HK, Stone SJG, Frantz AE. Human prolactin and growth hormone release during surgery and other conditions of stress. J Clin Endocrinol Metab 1972;35:840–851. 310 Harms PG, Langlier P, McCann SM. Modification of stress-induced prolactin release by dexamethasone or adrenalectomy. Endocrinology 1975;96:475–478. 311 Subramanian MG, Gala RR. The influence of adrenalectomy and of corticosterone administration on the ether-induced increase in plasma prolactin in ovariectomized estrogen-treated rats. Proc Soc Exp Biol Med 1978;157:415–417. 312 Van den Berghe G, de Zegher F, Bouillon R. Acute and prolonged critical illness as different neuroendocrine paradigms. J Clin Endocrinol Metab 1998; 83:1827–1834. 313 Gala RR. The physiology and mechanisms of the stress induced changes in prolactin secretion in the rat. Life Sci 1990;46:1407–1420. 314 Corenblum B, Taylor PJ. Mechanisms of control of prolactin release in response to apprehension stress and anesthesia-surgery success. Fertil Steril 1981;36:712–715. 315 Spiler IJ, Molitch ME. Lack of modulation of pituitary hormone stress response by neural pathways involving opiate receptors. J Clin Endocrinol Metab 1980;50:516–520. 316 Abe H, Engler D, Molitch ME et al. Vasoactive intestinal peptide is a physiological mediator of prolactin release in the rat. Endocrinology 1985;116: 1383–1390. 317 Kaji H, Chihara K, Kita T et al. Administration of antisera to vasoactive intestinal polypeptide and peptide histidine isoleucine attenuates etherinduced prolactin secretion in rats. Neuroendocrinology 1985;41:529–531. 318 Woolf PD, Lee LA, Leebaw WF. Hypoglycemia as a provocative test of prolactin release. Metabolism 1978;27:869–877. 319 Woolf PD, Lee LA, Leebaw W, Thompson D. Intracellular glucopenia causes prolactin release in man. J Clinic Endocrinol Metab 1977;45:377–382. 320 Whitaker MB, Corenblum B, Taylor PJ, Control of the hypoglycemia release of prolactin. Prog Reprod Biol 1980;6:77–82. 321 Chang FE, Dodds WG, Sullivan M, Kim MH. The acute effects of exercise on prolactin and growth hormone secretion: comparison between sedentary women and women runners with normal and abnormal menstrual cycles. J Clin Endocrinol Metab 1986;62:551–556. 322 Chang FE, Richards SR, Kim MH, Malarkey WB. Twenty-four hour prolactin profiles and prolactin responses to dopamine in long distance running women. J Clin Endocrinol Metab 1984;58:631–635. 323 Everett JW. Luteotrophic function of autografts of the rat hypophysis. Endocrinology 1954;54:685–690. 324 Pasteels JL. Administration d’extraits hypothalamiques a l’hypophyse de rat in vitro dans le but d’en controler la secretion de prolactine. C R Acad Sci Paris 1962;254:2664–2666. 325 Talwalker PK, Ratner A, Meites J. In vitro inhibition of pituitary prolactin synthesis and release by hypothalamic extract. Am J Physiol 1963;205:213– 218. 326 Barraclough CA, Sawyer CH. Induction of pseudopregnancy in the rat by reserpine and chlorpromazine. Endocrinology 1959;65:563–571. 327 Kanematsu S, Hilliard J, Sawyer CH. Effect of reserpine on pituitary prolactin content and its hypothalamic site of action in the rabbit. Acta Endocrin 44;1963:467–474. 328 van Maanen JH, Smelik PG. Induction of pseudopregnancy in rats following local depletion of monamines in the median eminence of the hypothalamus. Neuroendocrinology 1968;3:177–186. 329 Birge CA, Jacobs LS, Hammer CT, Daughaday WH. Catecholamine inhibition of prolactin secretion by isolated rat adenohypophyses. Endocrinology 1970;86:120–130.

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383 Enjalbert A, Moos F, Carbonell L et al. Prolactin inhibiting activity of dopamine-free subcellular fractions from rat mediobasal hypothalamus. Neuroendocrinology 1977;24:147–161. 384 Nikolics K, Mason AJ, Szonyi E et al. A prolactin-inhibiting factor within the precursor for human gonadotropin-releasing hormone. Nature 1985;316: 511–517. 385 Wormald PJ, Abrahamson MJ, Seeburg PH et al. Prolactin-inhibiting activity of GnRH associated peptide in cultured human pituitary cells. Clin Endocrinology 1989;30:149–155. 386 Chavali GB, Nagpal S, Majumdar SS et al. Helix-loop-helix motif in GnRH associated peptide is critical for negative regulation of prolactin secretion. J Mol Biol 1997;272:731–740. 387 Phillips HS, Nikolics K, Branton D, Seeburg PH. Immunocytochemical localization in rat brain of a prolactin release-inhibiting sequence of gonadotropin-releasing hormone prohormone. Nature 1985;316:542–545. 388 Clarke IJ, Cummins JT, Karsch FJ et al. GnRH-associated peptide (GAP) is cosecreted with GnRH into the hypophyseal portal blood of ovariectomized sheep. Biochem Biophys Res Comm 1987;143:665–671. 389 Adelman JP, Mason AJ, Hayflick JS, Seeburg PH. Isolation of the gene and hypothalamic cDNA for the common precursor of gonadotropin-releasing hormone and prolactin release-inhibiting factor in human and rat. Proc Natl Acad Sci USA 1986;83:179–183. 390 Ishibashi M, Yamaji T, Takaku F et al. Effect of GnRH-associated peptide on prolactin secretion from human lactotrope adenoma cells in culture. Acta Endocrinol 1987;116:81–84. 391 Thomas GB, Cummins JT, Doughton BW et al. Gonadotropin-releasing hormone associated peptide (GAP) and putative processed GAP peptides do not release luteinizing hormone or follicle-stimulating hormone or inhibit prolactin secretion in the sheep. Neuroendocrinology 1988;48:342–350. 392 Schally AV, Redding TW, Arimura A et al. Isolation of gamma-aminobutyric acid from pig hypothalami and demonstration of its prolactin releaseinhibiting (PIF) activity in vivo and in vitro. Endocrinology 1977;100:681–691. 393 Grandison L, Guidotti A. Gamma-aminobutyric acid receptor function in rat anterior pituitary: evidence for control of prolactin release. Endocrinology 1979;105:754–759. 394 Vincent SR, Hokfelt T, Wu JY. GABA neuron systems in the hypothalamus and the pituitary gland. Neuroendocrinology 1982;34:117–125. 395 Mulcahey JJ, Neill JD. Gamma-aminobutyric acid (GABA) levels in hypophyseal stalk plasma of rats. Life Sci 1982;32:453–456. 396 Lamberts SWJ, Macleod RM. Studies on the mechanism of the GABAmediated inhibition of prolactin secretion. Proc Soc Exptl Biol Med 1978;158: 10–13. 397 Grossman A, Delitala G, Yeo T, Besser GM. GABA and muscimol inhibit the release of prolactin from dispersed rat anterior pituitary cells. Neuroendocrinology 1981;32:145–149. 398 Loeffler JP, Kley N, Pittius CW et al. In vivo and in vitro studies of GABAergic inhibition of prolactin biosynthesis. Neuroendocrinology 1986;43:504–510. 399 Takahara J, Yunoki S, Yakushiji W et al. Stimulatory effects of gammahydroxybutyric acid on growth hormone and prolactin release in humans. J Clin Endocrinol Metab 1977;44:1014–1017. 400 Fioretti P, Melis GB, Paoletti AM et al. Gamma-Amino-b-hydroxy butyric acid stimulates prolactin and growth hormone release in normal women. J Clinic Endocrinol Metab 1978;47:1336–1340. 401 Tamminga CA, Neophytides A, Chase TN, Frohman LA. Stimulation of prolactin and growth hormone secretion by muscimol, a gamma aminobutyric acid agonist. J Clin Endocrinol Metab 1978;47:1348–1351. 402 Cavagnini F, Benetti G, Invitti C et al. Effect of gamma aminobutyric acid on growth hormone and prolactin secretion in man: influence of pimozide and domperidone. J Clin Endocrinol Metab 1980;51:789–797. 403 Melis GB, Fruzzetti F, Paoletti AM et al. Pharmacological activation of gaminobutyric acid-system blunts prolactin response to mechanical breast stimulation in puerperal women. J Clin Endocrinol Metab 1984;58:201–205. 404 Tashjian Jr AH, Barowsky NJ, Jensen DK. Thyrotropin releasing hormone: direct evidence for stimulation of prolactin production by pituitary cells in culture. Biochem Biophys Res Commun 1971;43:516–523. 405 Bowers CY, Friesen HG, Hwang P et al. Prolactin and thyrotropin release in man by synthetic pyroglutamyl-histidyl-prolinamide. Biochem Biophys Res Commun 1971;45:1033–1041. 406 Jacobs LS, Snyder PJ, Wilber JF et al. Increased serum prolactin after administration of synthetic thyrotropin releasing hormone (TRH) in man. J Clin Endocrinol Metab 1971;33:996–998. 407 Ashworth R, Hinkle PM. Thyrotropin-releasing hormone-induced intracellular calcium responses in individual rat lactotrophs and thyrotrophs. Endocrinology 1996;137:5205–5212.

408 Tashjian Jr AH, Heslop JP, Berridge MJ. Subsecond and second changes in inositol polyphosphates in GH4C1 cells induced by thyrotropin-releasing hormone. Biochem J 1987;243:305–308. 409 Morel G, Gourdji D, Grouselle D et al. Immunocytochemical evidence for in vivo internalization of thyroliberin into rat pituitary target cells. Neuroendocrinology 1985;41:312–320. 410 Le Dafniet M, Lefebvre P, Barret A et al. Normal and adenomatous human pituitaries secrete thyrotropin-releasing hormone in vitro: modulation by dopamine, haloperidol, and somatostatin. J Clin Endocrinol Metab 1990;71: 480–486. 411 Noel GL, Dimond RC, Wartofsky L et al. Studies of prolactin and TSH secretion by continuous infusion of small amounts of thyrotropin-releasing hormone (TRH). J Clin Endocrinol Metab 1974;39:6–17. 412 Koch Y, Goldhaber G, Fireman I et al. Suppression of prolactin and thyrotropin secretion in the rat by anti-serum to thyrotropin-releasing hormone. Endocrinology 1977;100:1476–1478. 413 Harris ARC, Christianson D, Smith MS et al. The physiological role of thyrotropin-releasing hormone in the regulation of thyroid-stimulating hormone and prolactin secretion in the rat. J Clin Invest 1978;61:441–448. 414 Sheward WJ, Fraser HM, Fink G. Effect of immunoneutralization of thyrotrophin-releasing hormone on the release of thyrotrophin and prolactin during suckling or in response to electrical stimulation of the hypothalamus in the anaesthetized rat. J Endocrinol 1985;106:113–119. 415 Horn AM, Fraser HM, Fink G. Effects of antiserum to thyrotrophin-releasing hormone on the concentrations of plasma prolactin, thyrotrophin and LH in the pro-oestrous rat. J Endocrinol 1985;104:205–209. 416 Fraser HM, McNeilly AS. Effect of chronic immunoneutralization of thyrotropin-releasing hormone on the hypothalamic–pituitary–thyroid axis, prolactin, and reproductive function in the ewe. Endocrinology 1982;111: 1964–1973. 417 Yamada M, Saga Y, Shibusawa N et al. Tertiary hypothyroidism and hyperglycemia in mice with targeted disruption of the thyrotropin-releasing hormone gene. Proc Natl Acad Sci USA 1997;94:10862–10867. 418 Blake CA. Stimulation of pituitary prolactin and TSH release in lactating and proestrus rats. Endocrinology 1974;94:503–508. 419 Burnet FR, Wakerley JB. Plasma concentrations of prolactin and thyrotrophin during suckling in urethane-anaesthetized rats. Endocrinology 1976;70:429– 437. 420 Riskind PN, Millard WJ, Martin JB. Evidence that thyrotropin-releasing hormone is not a major prolactin-releasing factor during suckling in the rat. Endocrinology 1984;115:312–316. 421 Gautvik KM, Tashjian Jr AH, Kourides IA et al. Thyrotropin-releasing hormone is not the sole physiologic mediator of prolactin release during suckling. N Engl J Med 1974;290:1162–1165. 422 Jeppsson S, Rannevik KO, Wide L. Influence of suckling and of suckling followed by TRH or LH-RH on plasma prolactin, TSH, GH and FSH. Acta Endocrinol 1976;82:246–253. 423 Segerson TP, Kauer J, Wolfe HC et al. Thyroid hormone regulates TRH biosynthesis in the paraventricular nucleus of the rat hypothalamus. Science 1987;238:78–80. 424 Rondeel JMM, DeGreef WJ, Schoot PVD et al. Effect of thyroid status and paraventricular area lesions on the release of thyrotropin-releasing hormone and catecholamines into hypophysial portal blood. Endocrinology 1988;123: 523–527. 425 DeLean A, Ferland L, Drouin J et al. Modulation of pituitary thyrotrophin releasing hormone receptor levels by estrogens and thyroid hormone. Endocrinology 1977;100:1505–1510. 426 Perrone MH, Hinkle PM. Regulation of pituitary receptors for thyrotropin releasing hormone by thyroid hormones. J Biol Chem 1978;253:5168– 5173. 427 Mori M, Yamada M. Thyroid hormones regulate the amount of thyrotrophinreleasing hormone in the hypothalamic median eminence of the rat. J Endocrinol 1987;114:443–448. 428 Snyder PJ, Jacobs LS, Utiger RD, Daughaday WH. Thyroid hormone inhibition of the prolactin response to thyrotropin-releasing hormone. J Clin Invest 1973;52:2324–2329. 429 Hermite ML, Robyn C, Golstein J et al. Prolactin and thyrotropin in thyroid diseases: lack of evidence for a physiological role of thyrotropin-releasing hormone in the regulation of prolactin secretion. Horm Metab Res 1974;6:1 90–195. 430 Yamaji T. Modulation of prolactin release by altered levels of thyroid hormones. Metabolism 1974;23:745–751. 431 Lechan RM, Wu P, Jackson IMD et al. Thyrotropin-releasing hormone precursor: Characterization in rat brain. Science 1986;231:159–161.

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in the control of growth hormone and prolactin release. Neuroendocrinology 1990;51:688–693. Sortino MA, Cronin MJ, Wise PM. Relaxin stimulates prolactin secretion from anterior pituitary cells. Endocrinology 1989;124:2013–2015. Camoratto AM, Grandison L. Platelet-activating factor stimulates prolactin release from dispersed rat anterior pituitary cells in vitro. Endocrinology 1989;124:1502–1506. Spangelo BL, Judd AM, Ross PC et al. Thymosin fraction 5 stimulates prolactin and growth hormone release from anterior pituitary cells in vitro. Endocrinology 1987;121:2035–2043. Waldhauser F, Lieberman HR, Lynch HJ et al. A pharmacological dose of melatonin increases PRL levels in males without altering those of GH, LH, FSH, TSH, testosterone or cortisol. Neuroendocrinology 1987;46:125–130. Khorram O, Mizunuma H, McCann SM. Effect of a-melanocyte-stimulating hormone on basal and stimulated release of prolactin: Evidence for dopaminergic mediation. Neuroendocrinology 1982;34:433–437. Khorram O, Bedran de Castro JC, McCann SM. Physiological role of amelanocyte-stimulating hormone in modulating the secretion of prolactin and luteinizing hormone in the female rat. Proc Natl Acad Sci USA 1984;81: 8004–8008. Larson GH, Koos RD, Sortino MA, Wise PM. Acute effect of basic fibroblast growth factor on secretion of prolactin as assessed by the reverse hemolytic plaque assay. Endocrinology 1990;126:927–932. Anderson ST, Curlewis JD. PACAP stimulates dopamine neuronal activity in the medial basal hypothalamus and inhibits prolactin. Brain Res 1998;790: 343–346. Knigge UP. Histaminergic regulation of prolactin secretion. Danish Med Bulletin 1990;37:109–124. Arakelian MC, Libertun C. H1 and H2 histamine receptor participation in the brain control of prolactin secretion in lactating rats. Endocrinology 1977; 100:890–895. Gibbs DM, Plotsky PM, De Greef WJ, Neill JD. Effect of histamine and acetylcholine on hypophysial stalk plasma dopamine and peripheral plasma prolactin levels. Life Sci 1979;24:2063–2070. Knigge U, Matzen S, Warberg J. Histaminergic stimulation of prolactin secretion mediated via H1- or H2-receptors: Dependence on routes of administration. Neuroendocrinology 1986;44:41–48. Knigge U, Dejgaard A, Wollesen F et al. Histamine regulation of prolactin secretion through H1- and H2-receptors. J Clin Endocrinol Metab 1982;55: 118–122. Carlson HE, Ippoliti AF. Cimetidine, an H2-antihistamine, stimulates prolactin secretion in man. J Clin Endocrinol Metab 1977;45:367–370. Pontiroli AE, Pellicciotta G, Alberetto M et al. Cimetidine and L-dopa in the control of prolactin secretion in man. Horm Metab Res 1979;11:257– 258. Sharpe PC, Melvin MA, Mills JG et al. Prolactin responses to histamine H2 receptor antagonists. Acta Endocrinol 1980;95:308–313. Masala A, Alagna S, Faedda R, Satta A, Rovasio PP. Prolactin secretion in man following acute and long-term cimetidine administration. Acta Endocrinol 1980;93:392–395. Samarthji L et al. Effect of clonidine on growth hormone, prolactin, luteinizing hormone, follicle-stimulating hormone, and thyroid-stimulating hormone in the serum of normal men. J Clin Endocrinol Metab 1975;41: 827–832. Terry C, Martin JB. Evidence of a-adrenergic regulation of episodic growth hormone and prolactin secretion in the undisturbed male rat. Endocrinology 1981;108:1869–1873. Lien EL, Morrison A, Kassarich J, Sullivan D. Alpha-2-adrenergic control of prolactin release. Neuroendocrinology 1986;44:184–189. Gold MS, Donabedian RK, Redmond DE Jr. Further evidence for alpha-2 adrenergic receptor mediated inhibition of prolactin secretion: The effect of yohimbine. Psychoneuroendocrinology 1979;3:253–260. Steiner J, Cassar J, Mashiter K et al. Effect of methyldopa on prolactin and growth hormone. Brit Med J 1976;1:1186–1188. Slater SL, Lipper S, Shiling DJ, Murphy DL. Elevation of plasma-prolactin by monoamine-oxidase inhibitors. Lancet 1977;2:275–276. Schlegel W, Wuarin F, Zbaren C, Zahnd GR. Lowering of cytosolic free CA2+ by carbachol, a muscarinic cholinergic agonist, in clonal pituitary cells (GH3 cells). Endocrinology 1985;117:976–981. Beach JE, Smallridge RC, Chiang PK, Fein HG. Reversal of inhibition of prolactin secretion in cultured pituitary cells by muscarinic antagonists. J Pharm Exper Ther 1988;246:548–552. Casanueva FF, Villanueva L, Diaz Y et al. Atropine selectively blocks GHRHinduced GH secretion without altering LH, FSH, TSH, PRL and

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ACTH/cortisol secretion elicited by their specific hypothalamic releasing factors. Clin Endocrinol 1986;25:319–323. Masala A, Alagna S, Devilla L et al. Muscarinic receptor blockade by pirenzepine: Effect on prolactin secretion in man. J Endocrinol Invest 1982;5:53–55. Mendelson WB, Sitaram N, Wyatt RJ, Gillin JC. Methscopolamine inhibition of sleep-related growth hormone secretion: Evidence for a cholinergic secretory mechanism. J Clin Invest 1978;61:1683–1690. Carmeliet P, Denef C. Immunocytochemical and pharmacological evidence for an intrinsic cholinomimetic system modulating prolactin and growth hormone release in rat pituitary. Endocrinology 1988;123:1128–1139. MacLeod RM, Smith MC, DeWitt GW. Hormonal properties of transplanted pituitary tumors and their relation to the pituitary gland. Endocrinology 1966;79:1149–1156. Chen CL, Minaguchi H, Meites J. Effects of transplanted pituitary tumors on host pituitary prolactin secretion. Proc Soc Exptl Biol Med 1967;126:317–320. Clemens JA, Meites J. Inhibition by hypothalamic prolactin implants of prolactin secretion, mammary growth and luteal function. Endocrinology 1968;82:878–881. Annunziato L, Moore KE. Prolactin in CSF selectively increases dopamine turnover in the median eminence. Life Sci 1978;22:2037–2042. Perkins NA, Westfall TC, Paul CV et al. Effect of prolactin on dopamine synthesis in medial basal hypothalamus: evidence for a short loop feedback. Brain Res 1979;160:431–444. Morgan WW, Herbert DC. Early responses of the dopaminergic tuberoinfundibular neurons to anterior pituitary homografts. Neuroendocrinology 1980;31:215–221. Selmanoff M. The lateral medial median eminence: distribution of dopamine, norepinephrine, and luteinizing hormone-releasing hormone and the effect of prolactin on catecholamine turnover. Endocrinology 1981;108:1716–1722. Bybee DE, Nakawatase C, Szabo M, Frohman LA. Inhibitory feedback effects of prolactin on its secretion involve central nervous system dopaminergic mediation. Neuroendocrinology 1983;36:27–32. Horseman ND, Zhao W, Montecino-Rodriguez E et al. Defective mammopoiesis, but normal hematopoiesis, in mice with a targeted disruption of the prolactin gene. EMBO J 1997;16:6926–6935. Shipman SL, Scheiber MD, Horseman ND. Pituitary hyperplasia in mice carrying a targeted disruption of the prolactin gene results from disordered dopaminergic control of lactotrophs. Program of the Meeting Endocrine Society, 1999, San Diego. (Abstract P2–491). Phelps CJ, Horseman ND. Hypophysiotropic dopaminergic neurons in prolactin-knockout mice. Program of the Meeting Endocrine Society, 1999, San Diego. (Abstract P2–492). Judd SJ. Autoregulation of prolactin secretion. Prog Reprod Biol 1980;6:87–91. Quigley ME, Yen SSC. Evidence for increased dopaminergic inhibition of secretion of thyroid-stimulating hormone in hyperprolactinemic patients with pituitary microadenoma. Am J Obstet Gynecol 1980;137:653–655. Scanlon MF, Rodriquez-Arnao MD, McGregor AM et al. Altered dopaminergic regulation of thyrotrophin release in patients with prolactinomas: comparison with other tests of hypothalamic-pituitary function. Clin Endocrinol 1981;14:133–143. Massara F, Camanni F, Martra M, Dolfin GC, Muller EE, Molinatti GM. Reciprocal pattern of the TSH and PRL responses to dopamine receptor blockade in women with physiological or pathological hyperprolactinemia. Clin Endocrinol 1983;18:103–110. Rodriguez-Arnao MD, Peters JR, Foord S et al. Exaggerated circadian variation in basal thyrotropin (TSH) and in the dopaminergic inhibition of TSH release in pathological hyperprolactinemia: evidence against a hypothalamic defect. J Clin Endocrinol Metab 1983;57:975–980. Holdaway IM, Evans MC, Sheehan A, Ibbertson HK. Low thyroxine levels in some hyperprolactinemic patients due to dopaminergic suppression of thyrotropin. J Clin Endocrinol Metab 1984;59:608–613. Ghigo E, Goffi S, Molinatti GM et al. Prolactin and TSH responses to both domperidone and TRH in normal and hyperprolactinaemic women after dopamine synthesis blockade. Clin Endocrinol 1985;23:155–160. Dieguez C, Peters JR, Page MD et al. Thyroid function in patients with hyperprolactinemia: relationship to dopaminergic inhibition of TSH release. Clin Endocrinol 1986;25:435–440. Winters SJ, Troen P. Altered pulsatile secretion of luteinizing hormone in hypogonadal men with hyperprolactinemia. Clin Endocrinol 1984;21:257–263. Sauder SE, Frager M, Case GD et al. Abnormal patterns of pulsatile luteinizing hormone secretion in women with hyperprolactinemia and amenorrhea: responses to bromocriptine. J Clin Endocrinol Metab 1984;59:941–948. Koizumi K, Aono T, Koike K, Kurachi K. Restoration of LH pulsatility in

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patients with prolactinomas after transsphenoidal surgery. Acta Endocrinol 1984;107:433–438. Wiebe RH, Handwerger S, Soules M. Episodic luteinizing hormone release in hyperprolactinemic women. Fertil Steril 1986;45:483–488. Molitch ME, Rebar RW, Barsano CP. Effect of human prolactin administration on gonadotropin and thyrotropin secretion in normal men. J Endocrinol Invest 1993;16:554–563. Sarkar DK. Evidence for prolactin feedback actions on hypothalamic oxytocin, vasoactive intestinal peptide and dopamine secretion. Neuroendocrinology 1989;49:520–524. Nicoll CS. Ontogeny and evolution of prolactin’s functions. Fed Proc 1980; 39:2563–2566. Whitworth NS. Lactation in humans. Psychoneuroendocrinology 1988;13: 171–188. Arden KC, Boutin J-M, Djiane J et al. The receptors for prolactin and growth hormone are localized in the same region of human chromosome 5. Cytogenet Cell Genet 1990;53:161–165. Hu Z-Z, Zhuang L, Meng J et al. The human prolactin receptor gene structure and alternative promoter utilization: the generic promoter hPIII and a novel human promoter hPn. J Clin Endocrinol Metab 1999;84:1153–1156. Boutin J-M, Edery M, Shirota M et al. Identification of a cDNA encoding a long form of prolactin receptor in human hepatoma and breast cancer cells. Mol Endocrinol 1989;3:1455–1461. Jolicoeur C, Boutin JM, Okamura H. Multiple regulators of prolactin receptor gene expression in rat liver. Mol Endocrinol 1989;3:895–900. Dunaif AE, Zimmerman EA, Friesen HG, Frantz AG. Intracellular localization of prolactin receptor and prolactin in the rat ovary by immunocytochemistry. Endocrinology 1982;110:1465–1472. Martin RH, Glass MR, Chapman C et al. Human a-lactalbumin and hormonal factors in pregnancy and lactation. Clin Endocrinol 1980;13:223– 230. Weinstein D, Ben-David M, Polishuk WZ. Serum prolactin and the suppression of lactation. Brit J of Obstet and Gynecol 1976;83:679–682. Ormandy CJ, Camus A, Barra J et al. Null mutation of the prolactin receptor gene produces multiple reproductive defects in the mouse. Genes Dev 1997;11:167–178. Thiede MA. The mRNA encoding a parathyroid hormone-like peptide is produced in mammary tissue in response to elevations in serum prolactin. Mol Endocrinol 1989;3:1443–1447. Khosla S, Johansen KL, Ory SJ et al. Parathyroid hormone-related peptide in lactation and in umbilical cord blood. Mayo Clin Proc 1990;65:1408–1414. Friedman S, Goldfien A. Breast secretions in normal women. Am J Obstet and Gynecol 1969;104:846–849. Lavric MV. Breast secretion in nulligravid women. Am J Obstet Gynecol 1972;112:1139–1140. Jones JR, Gentile GP. Incidence of galactorrhea in ovulatory and anovulatory females. Am J Obstet Gynecol 1975;45:13–14. Buckman MT, Peake GT. Incidence of galactorrhea. JAMA 1976;236:2747. Biller BJ, Boyd AE III, Molitch ME et al. Galactorrhea syndromes. In: Post KD, Jackson IMD, Reichlin S, eds. The Pituitary Adenoma, New York: Plenum Medical Book Co., 1980:65–90. Molitch ME, Reichlin S. Hyperprolactinemic disorders. Disease-a-Month 1982;28(9):1–58. Clément-Lacroix P, Ormandy C, Lepescheux L et al. Osteoblasts are a new target for prolactin: analysis of bone formation in prolactin receptor knockout mice. Endocrinology 1999;140:96–105. Steger RW, Chandrashekar V, Zhao W et al. Neuroendocrine and reproductive functions in male mice with targeted disruption of the prolactin gene. Endocrinology 1998;139:3691–3695. Kauppila A, Reinilä M, Martikainen H et al. Hypoprolactinemia and ovarian function. Fertil Steril 1988;49:437–441. Odell WD. Prolactin producing tumors (prolactinomas). In: Odell WD, Nelson D, eds. Pituitary Tumors. Mt. Kisco: Futura Publishing Co., Inc., 1984:159. Malarkey WB. Effects of hyperprolactinemia on other endocrine systems. In: Olefsky JM, Robbins RJ, eds. Prolactinomas. New York: Churchill Livingstone, 1986:21. Klibanski A, Beitins IZ, Merriam GR et al. Gonadotropin and prolactin pulsations in hyperprolactinemic women before and during bromocriptine therapy. J Clin Endocrinol Metab 1984;58:1141–1147. Ambrosi B, Giovine C, Nava C et al. Serum gonadotrophin pulsatile secretion in men with PRL-secreting and non-secreting pituitary tumours. Acta Endocrinol 1985;109:1–6. Fox SR, Hoefer MT, Bartke A, Smith MS. Suppression of pulsatile LH secretion, pituitary GnRH receptor content and pituitary responsiveness to

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GnRH by hyperprolactinemia in the male rat. Neuroendocrinology 1987;46: 350–359. Winters SJ, Loriaux DL. Suppression of plasma luteinizing hormone by prolactin in the male rat. Endocrinology 1978;102:864–868. Login IS, Krieg RJ, Kaiser DL et al. Unremitting suppression of the postovariectomy rise in plasma 1H by tumor-induced hyperprolactinemia. Neuroendocrinology 1982;35:327–332. Brar AK, McNeilly AS, Fink G. Effects of hyperprolactinaemia and testosterone on the release of LH-releasing hormone and the gonadotrophins in intact and castrated rats. J Endocrinol 1985;104:35–43. Soccia B, Schneider AB, Marut EL, Scommegna A. Pathological hyperprolactinemia suppresses hot flashes in menopausal women. J Clin Endocrinol Metab 1988;66:868–870. Sarkar DK, Yen SSC. Hyperprolactinemia decreases the luteinizing hormonereleasing hormone concentration in pituitary portal plasma: a possible role for b-endorphin as a mediator. Endocrinology 1985;116:2080–2084. Cheung CY. Prolactin suppresses luteinizing hormone secretion and pituitary responsiveness to luteinizing hormone-releasing hormone by a direct action at the anterior pituitary. Endocrinology 1983;113:632–638. Bohnet HG, Dahlen HG, Wuttke W, Schneider HPG. Hyperprolactinemia anovulatory syndrome. J Clin Endocrinol Metab 1975;42:132–137. Campenhout JV, Papas S, Blanchet P et al. Pituitary responses to synthetic luteinizing hormone-releasing hormone in thirty-four cases of amenorrhea or oligomenorrhea associated with galactorrhea. Am J Obstet Gynecol 1977;127: 723–728. Klibanski A, Beitins IZ, Zervas NT et al. a-subunit and gonadotropin responses to luteinizing hormone-releasing hormone in hyperprolactinemic women before and after bromocriptine. J Clin Endocrinol Metab 1983;56: 774–780. Marchetti B, Labrie F. Prolactin inhibits pituitary luteinizing hormonereleasing hormone receptors in the rat. Endocrinology 1982;111:1209–1216. Garcia A, Herbon L, Barkan A et al. Hyperprolactinemia inhibits gonadotropin-releasing hormone GnRH stimulation of the number of pituitary GnRH receptors. Endocrinology 1985;117:954–959. Duncan JA, Barkan A, Herbon L, Marshall JC. Regulation of pituitary gonadotropin-releasing hormone (GnRH) receptors by pulsatile GnRH in female rats: effects of estradiol and prolactin. Endocrinology 1986;118:320–327. Glass MR, Shaw RW, Butt WR et al. An abnormality of oestrogen feedback in amenorrhoea–galactorrhoea. British Medical J 1975;3:274–275. McNeilly AS, Glasier A, Jonassen J, Howie PW. Evidence for direct inhibition of ovarian function by prolactin. J Reprod Fert 1982;65:559–569. McNatty KP, Sawers RS, McNeilly AS. A possible role for prolactin in control of steroid secretion by the human Graafian follicle. Nature 1974;250:653–655. Feltus FA, Groner B, Melner MH. Stat5-mediated regulation of the human type II 3beta-hydroxysteroid dehydrogenase/delta5–delta4 isomerase gene: activation by prolactin. Mol Endocrinol 1999;13:1084–1093. Del Pozo E, Wyss H, Tolis G et al. Prolactin and deficient luteal function. Obstet Gynecol 1979;53:282–286. Schulz KD, Geiger W, Del Pozo E, Künzig HJ. Pattern of sexual steroids, prolactin, and gonadotropic hormones during prolactin inhibition in normally cycling women. Am J Obstet Gynecol 1978;132:561–566. Mühlenstedt D, Bohnet HG, Hanker JP, Schneider HPG. Short luteal phase and prolactin. Int J Fertil 1978;23:213–218. Bahamondes L, Saboya W, Tambascia M, Trevisan M. Galactorrhea, infertility, and short luteal phases in hyperprolactinemic women: early state of amenorrhea-galactorrhea? Fertil Steril 1979;32:476–477. McNatty KP. Relationship between plasma prolactin and the endocrine microenvironment of the developing human antral follicle. Fertil Steril 1979;32:433–438. Demura R, Ono M, Demura H et al. Prolactin directly inhibits basal as well as gonadotropin-stimulated secretion of progesterone and 17b-estradiol in the human ovary. J Clin Endocrinol Metab 1982;54:1246–1250. Veldhuis JD, Hammond JM. Oestrogens regulate divergent effects of prolactin in the ovary. Nature 1980;284:262–264. Dorrington JH, Gore-Langton RE. Antigonadal action of prolactin: Further studies on the mechanism of inhibition of follicle-stimulating hormoneinduced aromatase activity in rat granulosa cell cultures. Endocrinology 1982;110:1701–1707. Krasnow JS, Hickey GJ, Richards JS. Regulation of aromatase mRNA and estradiol biosynthesis in rat ovarian granulosa and luteal cells by prolactin. Mol Endocrinol 1990;4:13–21. Magoffin DA, Erickson GF. Prolactin inhibition of luteinizing hormonestimulated androgen synthesis in ovarian interstitial cells cultured in defined medium: Mechanism of action. Endocrinology 1982;111:2001–2007.

722 Fraser IS, Markham R, Shearman RP. Plasma prolactin levels and ovarian responsiveness to exogenous gonadotropins. Obstet Gynecol 1978;51:548–551. 723 McGarrigle HHG, Sarris S, Little V et al. Induction of ovulation with clomiphene and human chorionic gonadotrophin in women with hyperprolactinaemic amenorrhoea. Brit J Obstet Gynecol 1978;85:692–697. 724 Duignant NM. Polycystic ovarian disease. Brit J Obstet Gynaecol 1976;83: 593–602. 725 Futterweit W, Krieger DT. Pituitary tumors associated with hyperprolactinemia and polycystic ovarian disease. Fertil Steril 1979;31:608–613. 726 Del Pozo E, Falaschi P. Prolactin and cyclicity in polycystic ovary syndrome. Prog Reprod Biol 1980;6:252–259. 727 Buvat J, Mourot CS, Fourlinnie JC et al. Androgens and prolactin levels in hirsute women with either polycystic ovaries or “borderline ovaries”. Fertil Steril 1982;38:695–700. 728 Corenblum B, Taylor PJ. The hyperprolactinemic polycystic ovary syndrome may not be a distinct entity. Fertil Steril 1982;38:549–552. 729 Skrabanek P, McDonald D, De Valera E et al. Plasma prolactin in amenorrhoea, infertility, and other disorders: a retrospective study of 608 patients. Irish J Med Sci 1980;149:236–245. 730 Kemmann E, Jones JR. Hyperprolactinemia and primary amenorrhea. Obstet Gynecol 1979;54:692–694. 731 Mashchak CA, Kletzky OA, Davajan V, Mishell DR. Clinical and laboratory evaluation of patients with primary amenorrhea. Obstet Gynecol 1981;57: 715–721. 732 Pepperell RJ, Aust F. Prolactin and reproduction. Fertil Steril 1981;35:267–274. 733 Seppala M, Hirvonen E, Ranta T. Hyperprolactinaemia and luteal insufficiency. Lancet 1976;1:229–230. 734 Kredentser JV, Hoskins CF, Scott JZ. Hyperprolactinemia—a significant factor in female infertility. Am J Obstet Gynecol 1981;139:264–267. 735 Huang K-E, Bongiglio TA, Muschler EK. Transient hyperprolactinemia in infertile women with luteal phase deficiency. Obstet Gynecol 1991;78:651–655. 736 Fava GA, Fava M, Kellner R et al. Depression, hostility and anxiety in hyperprolactinemic amenorrhea. Psychother Psychosom 1981;36:122–128. 737 Post KD, Biller BJ, Adelman LS et al. Selective transsphenoidal adenomectomy in women with galactorrhea–amenorrhea. JAMA 1979;242:158–162. 738 Bartke A. Effects of prolactin on spermatogenesis in hypophysectomized mice. Endocrinology 1971;49:311–316. 739 Hafiez AA, Bartke A, Lloyd CW. The role of prolactin in the regulation of testis function: the synergistic effects of prolactin and luteinizing hormone on the incorporation of [1-14C] acetate into testosterone and cholesterol by testes from hypophysectomized rats in vitro. Endocrinology 1972;53:223–230. 740 Aragona C, Bohnet HG, Friesen HG. Localization of prolactin binding in prostate and testis: the role of serum prolactin concentration on the testicular LH receptor. Acta Endocrinol 1977;84:402–409. 741 Musto N, Hafiez AA, Bartke A. Prolactin increases 17b-hydroxysteroid dehydrogenase activity in the testis. Endocrinology 1972;95:1106–1108. 742 Hafriez AA, Philpott JE, Bartke A. The role of prolactin in the regulation of testicular function: the effect of prolactin and luteinizing hormone on 3bhydroxysteroid dehydrogenase activity in the testes of mice and rats. Endocrinology 1971;50:619–623. 743 Oseko F, Endo J, Nakano A et al. Effects of chronic bromocriptine-induced hypoprolactinemia on plasma testosterone responses to human chorionic gonadotropin stimulation in normal men. Fertil Steril 1991;55:355–357. 744 Sheth AR, Mugatwala PP, Shah GV, Rao SS. Occurrence of prolactin in human semen. Fertil Steril 1975;26:905–907. 745 Shah GV, Desai RB, Sheth AR. Effect of prolactin of metabolism of human spermatozoa. Fertil Steril 1976;27:1292–1294. 746 Carter JN, Tyson JE, Tolis G et al. Prolactin-secreting tumors and hypogonadism in 22 men. New Engl J Med 1978;299:847–852. 747 Nagulesparen M, Ang V, Jenkins JS. Bromocriptine treatment of males with pituitary tumours, hyperprolactinaemia, and hypogonadism. Clin Endocrinol 1978;9:73–79. 748 Franks S, Jacobs HS, Martin N, Nabarro JDN. Hyperprolactinaemia and impotence. Clin Endocrinol 1978;8:277–287. 749 Goodman RH, Molitch ME, Post KD, Jackson IMD. Prolactin-secreting adenomas in the male. In: Post KD, Jackson IMD, Reichlin S, eds. The Pituitary Adenoma. New York: Plenum Medical Book Co., 1980:91–108. 750 Ambrosi B, Gaggini M, Travaglini P et al. Hypothalamic–pituitary–testicular function in men with PRL-secreting tumors. J Endocrinol Invest 1981;4: 309–315. 751 Grisoli F, Vincentelli F, Jaquet P et al. Prolactin secreting adenoma in 22 men. Surg Neurol 1980;13:241–247. 752 Spark RF, O’Reilly G, Wills CA et al. Hyperprolactinaemia in males with and without pituitary macroadenomas. Lancet 1982;2:129–131.

Chapter 4 753 Prescott RWG, Kendall-Taylor P, Hall K et al. Hyperprolactinaemia in men— response to bromocriptine therapy. Lancet 1982;1:245–248. 754 Perryman RL, Thorner MO. The effects of hyperprolactinemia on sexual and reproductive function in men. J Androl 1981;5:233–242. 755 Dupuy M, Derome PJ, Peillon F et al. L’adenome a prolactine chez l’homme. Sem Hôp Paris 1984;60:2943–2954. 756 Heshmati HM, Turpin G, Nahoul K et al. Testicular response to human chorionic gonadotrophin in chronic hyperprolactineamia. Acta Endocrinol 1985;108:565–569. 757 Luboshitzky R, Rosen E, Trestian S, Spitz M. Hyperprolactinaemia and hypogonadism in men: Response to exogenous gonadotrophins. Clin Endocrinol 1979;11:217–223. 758 Murray FT, Cameron DF, Ketchum C. Return of gonadal function in men with prolactin-secreting pituitary tumors. J Clin Endocrinol Metab 1984;59: 79–85. 759 Molitch ME, Elton RL, Blackwell RE et al. Bromocriptine as primary therapy for prolactin-secreting macroadenomas: results of a prospective multicenter study. J Clin Endocrinol Metab 1985;60:698–705. 760 Magrini G, Ebiner JR, Burckhardt P, Felber JP. Study on the relationship between plasma prolactin levels and androgen metabolism in man. J Clin Endocrinol Metab 1976;43:944–947. 761 Cameron DF, Murray FT, Drylie DD. Ultrastructural lesions in testes from hyperprolactinemic men. J Androl 1984;5:283–293. 762 Schwartz MF, Bauman JE, Masters WH. Hyperprolactinemia and sexual disorders in men. Biol Psych 1982;17:861–876. 763 Leonard MP, Nickel CJ, Morales A. Hyperprolactinemia and impotence: why, when and how to investigate. J Urol 1989;142:992–994. 764 Buvat J, Lemaire A. Endocrine screening in 1,022 men with erectile dysfunction: clinical significance and cost-effective strategy. J Urol 1997;158: 1764–1767. 765 Segal S, Polishuk WZ, Ben-David M. Hyperprolactinemic male infertility. Fertil Steril 1976;27:1425–1427. 766 Hargreave JB, Kyle KF, Kelly AM, England P. Prolactin and gonadotrophins in 208 men presenting with infertility. Brit J Urology 1979;49:747–750. 767 Rjosk HK, Schill WB. Serum prolactin in male infertility. Andrologia 1979;11: 297–304. 768 Hargreave TB, Richmond JD, Liakatas J et al. Searching for the infertile man with hyperprolactinemia. Fertil Steril 1981;36(5):630–632. 769 Carter JN, Tyson JE, Warne GL et al. Adrenocortical function in hyperprolactinemic women. J Clin Endocrinol Metab 1977;45:973–980. 770 Vermeulen A, Ando S. Prolactin and adrenal androgen secretion. Clin Endocrinol 1978;8:295–303. 771 Lobo RA, Kletzky OA, Kaptein EM, Goebelsmann U. Prolactin modulation of dehydroepiandrosterone sulfate secretion. Am J Obstet Gynecol 1980;138: 632–636. 772 Parker LN, Change S, Odell WD. Adrenal androgens in patients with chronic marked elevation of prolactin. Clin Endocrinol 1978;8:1–5. 773 Belisle S. Adrenal androgen production in hyperprolactinemic states. Fertil Steril 1980;33:396–400. 774 Glickman SP, Rosenfield RL, Bergenstal RM, Helke J. Multiple androgenic abnormalities, including elevated free testosterone, in hyperprolactinemic women. J Clin Endocrinol Metab 1982;55:251–257. 775 Vermeulen A, Andó, Verdonck L. Prolactinomas, testosterone-binding globulin, and androgen metabolism. J Clin Endocrinol Metab 1982;54:409–412. 776 Lobo RA, Kletzky OA. Normalization of androgen and sex hormonebinding globulin levels after treatment of hyperprolactinemia. J Clin Endocrinol Metab 1983;56:562–566. 777 Pahuja DN, DeLuca HF. Stimulation of intestinal calcium transport and bone calcium mobilization by prolactin in vitamin D-deficient rats. Science 1981;214:1038–1039. 778 Spanos E, Brown DJ, Stevenson JC, Maclntyre IM. Stimulation of 1,25dihydroxycholecalciferol production by prolactin and related peptides in intact renal cell preparations in vitro. Biochem Biophys Acta 1981;672:7–15. 779 Kumar R, Abboud CF, Riggs BL. The effect of elevated prolactin levels on plasma 1,25-dihydroxvitamin D and intestinal absorption of calcium. Mayo Clin Proc 1980;55:51–53. 780 Wark JD, Tashjian AH Jr. Vitamin D stimulates prolactin synthesis by GH4C1 cells incubated in chemically defined medium. Endocrinology 1982;111: 1755–1757. 781 Wark JD, Tashjian AH Jr. Regulation of prolactin mRNA by 1,25dihydroxyvitamin D3in GH4C1 cells. J Biol Chem 1983;258:12118–12121. 782 Tornquist K, Allardt-Lamberg C. Systemic effects of 1,25-dihydroxyvitamin D3 on the pituitary-hypothalamic axis in rats. Acta Endocrinol 1987;115:225–228. 783 Tornquist K, Tashjian AH JR. Dual actions of 1,25-dihydroxycholecalciferol

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on intracellular Ca2+ in GH4C1 cells: evidence for effects on voltage-operated Ca2+ channels and Na+/Ca2+ exchange. Endocrinology 1989;124(6):2765–2776. Klibanski A, Neer RM, Beitins IZ et al. Decreased bone density in hyperprolactinemic women. N Engl J Med 1980;303:1511–1514. Schlechte JA, Sherman B, Martin R. Bone density in amenorrheic women with and without hyperprolactinemia. J Clin Endocrinol Metab 1983;56:1120– 1123. Koppelman MCS, Kurtz DW, Morrish KA et al. Vertebral body bone mineral content in hyperprolactinemic women. J Clin Endocrinol Metab 1984;59:1050– 1053. Schlechte J, El-Khoury G, Kathol M, Walkner L. Forearm and vertebral bone mineral in treated and untreated hyperprolactinemic amenorrhea. J Clin Endocrinol Metab 1987;64:1021–1026. Klibanski A, Greenspan SL. Increase in bone mass after treatment of hyperprolactinemic amenorrhea. New Engl J Med 1986;315(9):542–546. Caraceni MP, Gorhi E, Ortolani S et al. Increased forearm bone mineral content after bromocriptine treatment in hyperprolactinemia. Calcif Tissue Int 1985;37:687–689. Ciccarelli E, Savino L, Carlevatto V et al. Vertebral bone density in nonamenorrhoeic hyperprolactinaemic women. Clin Endocrinol 1988;28:1–6. Klibanski A, Biller BMK, Rosenthal DI et al. Effects of prolactin and estrogen deficiency in amenorrheic bone loss. J Clin Endocrinol Metab 1988;67:124– 130. Greenspan SL, Neer RM, Ridgway EC, Klibanski A. Osteoporosis in men with hyperprolactinemic hypogonadism. Ann Intern Med 1986;104:777–782. Greenspan SL, Oppenheim DS, Klibanski A. Importance of gonadal steroids to bone mass in men with hyperprolactinemic hypogonadism. Ann Intern Med 1989;110:526–531. Tesone M, Filho-Oliverira RM, Charreau EH. Prolactin binding in rat Langerhans islets. J Receptor Res 1980;1:355–372. Landgraf R, Leurs-Landgraf MMC, Weissmann A et al. Prolactin: a diabetogenic hormone. Diabetologia 1977;13:99–104. Gustafson AB, Banasiak MF, Kalkhoff RK et al. Correlation of hyperprolactinemia with altered plasma insulin and glucagon: Similarity to effects of late human pregnancy. J Clin Endocrinol Metab 1980;51:242–246. Schernthaner G, Prager R, Punzengruber C, Luger A. Severe hyperprolactinaemia is associated with decreased insulin binding in vitro and insulin resistance in vivo. Diabetologia 1985;28:138–142. Scobie IN, Kesson CM, Ratcliffe JG, MacCuish AC. The effects of prolonged bromocriptine administration on PRL secretion, GH and glycaemic control in stable insulin-dependent diabetes mellitus. Clin Endocrinol 1983;18:179– 185. Loretz CA, Bern HA. Prolactin and osmoregulation in vertebrates. Neuroendocrinology 1982;35:292–304. Adler RA, Noel GL, Wartofsky L, Frantz AG. Failure of oral water loading and intravenous hypotonic saline to suppress plasma prolactin in man. J Clin Endocrinol 1975;41:383–389. Baumann G, Marynick SP, Winters SJ, Loriaux DL. The effect of osmotic stimuli on prolactin secretion and renal water excretion in normal man and in chronic hyperprolactinemia. J Clin Endocrinol Metob 1977;44:199–202. Baumann G, Loriaux L. Failure of endogenous prolactin to alter renal salt and water excretion and adrenal function in man. J Clin Endocrinol Metab 1976;43:643–649. Sowers JR, Hershman JM, Skowsky WR et al. Osmotic control of the release of prolactin and thyrotropin in euthyroid subjects and patients with pituitary tumors. Metabolism 1977;26:187–192. Nagy E, Berczi I. Immunodeficiency in hypophysectomized rats. Acta Endocrinol 1978;89:530–537. Nagy E, Berczi I, Friesen HG. Regulation of immunity in rats by lactogenic and growth hormones. Acta Endocrinol 1983;102:351–357. Bernton EW, Meltzer MS, Holaday JW. Suppression of macrophage activation and t-lymphocyte function in hypoprolactinemic mice. Science 1988;239: 401–404. Hartman DP, Holaday JW, Bernton EW. Inhibition of lymphocyte proliferation by antibodies to prolactin. FASEB 1989;3:2194–2202. Montgomery DW, Zukoski CF, Shah GN. Conconavalin A-stimulated murine splenocytes produce a factor with prolactin-like bioactivity and immunorreactivity. J Biochem and Biophy 1987;145(2):692–698. Hiestand PC, Mekler P, Nordmann R. Prolactin as a modulator of lymphocyte responsiveness provides a possible mechanism of action for cyclosporine. Proc Natl Acad Sci USA 1986;83:2599–2603. Sabharwal P, Glaser R, Lafuse W et al. Prolactin synthesized and secreted by human peripheral blood mononuclear cells: an autocrine growth factor for lymphoproliferation. Proc Natl Acad Sci 1992;89:7713–7716.

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811 Pelligrini K, Lebrun J-J, Ali S, Kelly PA. Expression of prolactin and its receptor in human lymphoid cells. Mol Endocrinol 1992;6:1023–1031. 812 Wu H, Devi R, Malarkey WB. Expression and localization of prolactin messenger ribonucleic acid in the human immune system. Endocrinology 1996;137:349–353. 813 Matera L. Endocrine, paracrine and autocrine actions of prolactin on immune cells. Life Sci 1996;59:599–614. 814 Azad N, Agrawal L, Emanuele MA et al. Neuroimmunoendocrinology. Am J Reprod Immunol 1991;26:160–172. 815 Smith EM. Hormonal activities of cytokines. In: Blalock JE ed. Neuroimmunoendocrinology, 2nd edn. Basel: S. Karger 1992:154–169. 816 Mandrup-Poulsen T, Nerup J, Reimers JI et al. Cytokines and the endocrine system. I. The immunoendocrine network. Eur J Endocrinol 1995;133:660–671. 817 Di Carlo R, Meli R, Galdiero M et al. Prolactin protection against lethal effects of salmonella typhimurium. Life Sci 1993;53:981–989. 818 McMurray R, Keisler D, Kanuckel K et al. Prolactin influences autoimmune disease activity in the female B/W mouse. J Immunol 1991;147:3780–3787. 819 Harris RD, Kay NE, Seljeskog EL et al. Prolactin suppression of leukocyte chemotaxis in vitro. J Neurosurg 1979;50:462–465. 820 Fornari MC, Palacios MF, Diez RA, Intebi AD. Decreased chemotaxis of neutrophils in acromegaly and hyperprolactinemia. Eur J Endocrinol 1994;130:463–468. 821 Sabharwal P, Zwilling B, Glaser R, Malarkey WB. Cellular immunity in patients with acromegaly and prolactinomas. Prog Neuro Endocrin Immunology 1992;5:120–125. 822 Vidaller A, Guadarrama F, Llorente L et al. Hyperprolactinemia inhibits natural killer (NK) cell function in vivo and its bromocriptine treatment not only corrects it but makes it more efficient. J Clin Immunol 1992;12:210– 215. 823 Nagy E, Chalmers IM, Baragar FD et al. Prolactin deficiency in rheumatoid arthritis. J Rheumatol 1991;18:1662–1668. 824 Lavalle C, Loyo E, Paniagua R et al. Correlation study between prolactin and androgens in male patients with systemic lupus erythematosus. J Rheumatol 1987;14:268–272. 825 Polomeev M, Prokaeva T, Nassonova V et al. Prolactin levels in men with SLE and RA. J Rheumatol 1990;17:1569–1570. 826 Jara LJ, Gomez-Sanchez C, Silveira LH et al. Hyperprolactinemia in systemic lupus erythematosus: association with disease activity. Am J Med Sci 1992;303:222–226. 827 Pauzner R, Urowitz MB, Gladman DD, Gough JM. Prolactin in systemic lupus erythematosus. J Rheumatol 1994;21:2064–2067. 828 Neidhart M. Elevated serum prolactin or elevated Prolactin/cortisol ratio are associated with autoimmune processes in systemic lupus erythematosus and other connective tissue diseases. J Rheumatol 1996;23:476–481. 829 El-Garf A, Salah S, Shaarawy M et al. Prolactin hormone in juvenile systemic lupus erythematosus: a possible relationship to disease activity and CNS manifestations. J Rheumatol 1996;23:374–377. 830 Rovensky J, Juránková E, Rauová L et al. Relationship between endocrine, immune and clinical variables in patients with systemic lupus erythematosus. J Rheumatol 1997;24:2330–2334. 831 Mok CC, Lau CS, Lee KW, Wong RWS. Hyperprolactinemia in males with systemic lupus erythematosus. J Rheumatol 1998;25:2357–2373. 832 Funauchi M, Ikoma S, Enomoto H et al. Prolactin modulates the disease activity of systemic lupus erythematosus accompanied by prolactinoma. Clin Extpl Rheumatol 1998;16:479–482. 833 Jara LJ, Silveira LH, Cuellar ML et al. Hyper-prolactinemia in Reiter’s syndrome. J Rheumatol 1994;21:1292–1297. 834 Bravo G, Zazueta B, Lavalle C. An acute remission of Reiter’s syndrome in male patients treated with bromocriptine. J Rheumatol 1992;19:747–750. 835 Giasuddin ASM, El-Sherif AI, El-Ojali SI. Prolactin: does it have a role in the pathogenesis of psoriasis? Dermatology 1998;197:119–122. 836 Murphy WJ, Rui H, Longo DL. Effects of growth hormone and prolactin immune development and function. Life Sci 1995;57:1–14. 837 Josimovich JB, Lavenhar MA, Devanesan MM. Heterogeneous distribution of serum prolactin values in apparently healthy young women, and the effects of oral contraceptive medication. Fertil Steril 1987;47:785–791. 838 Miyai K, Ichihara K, Kondon K, Mori S. Asymptomatic hyperprolactinaemia and prolactinoma in the general population-mass screening by paired assays of serum prolactin. Clin Endocrinol 1986;25:549–554. 839 Miyake A, Ikegami M, Chen C-F. Mass screening for hyperprolactinemia and prolactinoma in men. J Endocrinol Invest 1988;11:383–384. 840 Jeffcoate SL. Diagnosis of hyperprolactinemia. Lancet 1978;2:1245–1248. 841 Kauppila A, Chatelain P, Kirkinen P. Isolated prolactin deficiency in a woman with puerperal alactogenesis. J Clin Endocrinol Metab 1987;64:309–312.

842 Shahmanesh M, Ali Z, Pourmand M et al. Pituitary function tests in Sheehan’s syndrome. Clin Endocrinol 1980;12:303–311. 843 Carlson HE, Brickman AS, Bottazzo GF. Prolactin deficiency in pseudohypoparathyroidism. N Engl J Med 1977;296:140–144. 844 De Rivera JL, Lal S, Ettigi P, Hontela S. Effect of acute and chronic neuroleptic therapy on serum prolactin levels in men and women of different age groups. Clin Endocrinology 1976;5:273–282. 845 Spitzer M, Sajjad R, Benjamin F. Pattern of development of hyperprolactinemia after initiation of haloperidol therapy. Obstet Gynecol 1998;91:693–695. 846 Kleinberg DL, Davis JM, de Coster R et al. Prolactin levels and adverse events in patients treated with risperidone. J Clin Psychopharmacol 1999;19:57–61. 847 Meltzer HY, Fang VS, Tricou BJ, Robertson A. Effect of antidepressants on neuroendocrine axis in humans. In: Costa E, Racagni G, eds. Typical and Atypical Antidepressants: Clinical Practice. New York: Raven Press, 1982:303–316. 848 Cowen PJ, Sargent PA. Changes in plasma prolactin during SSRI treatment: evidence for a delayed increase in 5-HT neurotransmission. J Psychopharmacol 1997;11:345–348. 849 Amsterdam JD, Garcia-España F, Goodman D et al. Breast enlargement during chronic antidepressant therapy. J Affect Dis 1997;46:151–156. 850 Mendelson JH, Mello NK, Teoh SK. Cocaine effects on pulsatile secretion of anterior pituitary, gonadal, and adrenal hormones. J Clin Endocrin Metab 1989;69:1256–1260. 851 Lee PA, Kelly MR, Wallin JD. Increased prolactin levels during reserpine treatment of hypertensive patients. JAMA 1976;235:2316–2317. 852 Barbarino A, De Marinis L. Calcium antagonists and hormone release. II. Effects of verapamil on basal, gonadotropin-releasing hormone-and thyrotropin-releasing hormone-induced pituitary hormone release in normal subjects. J Clin Endocrinol Metab 1980;51:749–753. 853 Maestri E, Camellini L, Rossi G, Bordonali G. Effects of five days verapamil administration on serum GH and PRL levels. Horm Metabol Res 1985;17: 482. 854 Romeo JH, Dombrowski R, Kwak YS et al. Hyperprolactinaemia and verapamil: prevalence and potential association with hypogonadism in men. Clin Endocrinol 1996;45:571–575. 854a Hutchinson J, Murphy M, Harries R et al. Galactorrhea and hyperprolactinemia associated with protease inhibitors. Lancet 2000;356:1003–1004. 855 Miyabo S, Hisada T, Asato T et al. Growth hormone and cortisol responses to psychological stress: comparison of normal and neurotic subjects. J Clin Endocrinol Metab 1976;42:1158–1162. 856 Yen SSC, Rebar RW, Quesenberry W. Pituitary function in pseudocyesis. J Clin Endocrinol Metab 1976;43:132–136. 857 Lim VS, Kathpalia SC, Frohman LA. Hyperprolactinemia and impaired pituitary response to suppression and stimulation in chronic renal failure: reversal after transplantation. J Clin Endocrinol Metab 1979;48:101–107. 858 Gomez F, Reyes FI, Faiman C. Nonpuerperal galactorrhea and hyperprolactinemia. Am J Med 1977;62:648–660. 859 Hou SH, Grossman S, Molitch ME. Hyperprolactinemia in patients with renal insufficiency and chronic renal failure requiring hemodialysis or chronic ambulatory peritoneal dialysis. Am J Kidney Dis 1985;6:245–249. 860 Morgan MY, Jakobovits AW, Gore MB et al. Serum prolactin in liver disease and its relationship to gynecomastia. Gut 1978;19:170–174. 861 Van Thiel DH, McClain CJ, Elson MK et al. Evidence for autonomous secretion of prolactin in some alcoholic men with cirrhosis and gynecomastia. Metabolism 1978;27:1778–1784. 862 Majumdar SK. Serum prolactin in chronic alcoholics. The Practitioner 1979;222:693–695. 863 Nunziata V, Ceparano G, Mazzacca G, Budillon G. Prolactin secretion in nonalcoholic liver cirrhosis. Digestion 1978;18:157–161. 864 Panerai AE, Salerno F, Manneschi M et al. Growth hormone and prolactin responses to thyrotropin-releasing hormone in patients with severe liver disease. J Clin Endocrinol Metab 1977;45:134–140. 865 Corenblum B, Shaffer EA. Hyperprolactinemia in hepatic encephalopathy may result from impaired central dopaminergic neurotransmission. Horm Metab Res 1989;21:675–677. 866 Välimäki M, Pelkonen R, Härkonen M, Tuomala P. Pituitary-gonadal hormones and adrenal androgens in non-cirrhotic female alcoholics after cessation of alcohol intake. Eur J Clin Invest 1990;20:177–181. 867 Honbo KS, Herle AJV, Kellett KA. Serum prolactin levels in untreated primary hypothyroidism. Am J Med 1978;64:782–787. 868 Smallridge RC. Thyrotropin-secreting pituitary tumors. Endocrinol Metab Clinics N Amer 1987;16:765–792. 869 Ben-David M, Danon A, Benveniste R et al. Results of radioimmunoassays of

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871 872

873 874

875 876 877 878 879 880 881 882 883

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rat pituitary and serum prolactin after adrenalectomy and perphenazine treatment in rats. J Endocrinol 1972;50:559–606. Refetoff S, Block MB, Ehrlich EN, Friesen GH. Chiari-Frommel syndrome in a patient with primary adrenocortical insufficiency. N Engl J Med 1972; 287:1326–1328. Lever EG, McKerron CG. Auto-immune Addison’s disease associated with hyperprolactinaemia. Clin Endocrinology 1984;21:451–457. Stryker TD, Molitch ME. Reversible hyperthyrotropinemia, hyperthyroxinemia and hyperprolactinemia due to adrenal insufficiency. Am J Med 1985;79:271–276. Kelver ME, Nagamani M. Hyperprolactinemia in primary adrenocortical insufficiency. Fertil Steril 1985;44:422–425. Shibutani Y. Case report: prolactin dynamics in a patient with isolated ACTH deficiency accompanied by hyperprolactinemia. Am J Med Sci 1988;295:140– 143. Boyd AE III, Spare S, Bower B, Reichlin S. Neurogenic galactorrhea–amenorrhea. J Clin Endocrinol Metab 1978;47:1374–1377. Morley JE, Hodgkinson DH, Kalk WJ. Galactorrhea and hyperprolactinemia associated with chest wall injury. J Clin Endocrinol Metab 1977;45:931–935. Herman VS, Kalk WJ. Neurogenic prolactin release: effects of mastectomy and thoracotomy. Prog Reprod Biol 1980;6:83–86. Wang Y-H, Huang T-S, Lien I-N. Hormone changes in men with spinal cord injuries. Am J Phys Med Rehabil 1992;71:328–322. Molitch ME, Schwartz S, Mukherji B. Is prolactin secreted ectopically? Am J Med 1981;70:803–807. Hsu C-T, Yu M-H, Lee C-YG et al. Ectopic production of prolactin in uterine cervical carcinoma. Gynecol Oncol 1992;44:166–171. Stanisic TH, Donova J. Prolactin secreting renal cell carcinoma. J Urol 1986;136:85–86. Hoffman WH, Gala RR, Kovacs K, Subramanian MG. Ectopic prolactin secretion from a gonadoblastoma. Cancer 1987;60:2690–2695. Palmer PE, Bogojavlensky S, Bhan AK, Scully RE. Prolactinoma in wall of ovarian dermoid cyst with hyperprolactinemia. Obstet Gynecol 1990;75: 540–543. Kallenberg GA, Pesce CM, Norman B. Ectopic hyperprolactinemia resulting from an ovarian teratoma. JAMA 1990;263:2472–2474. Molitch ME, Reichlin S. Hypothalamic hyperprolactinemia: neuroendocrine regulation of prolactin secretion in patients with lesions of the hypothalamus and pituitary stalk. In: MacLeod RM, Thorner MO, Scapagnini U, eds. Prolactin. Basic and Clinical Correlates. Padova: Liviana Press 1985:709–719. Snyder PJ, Jacobs LS, Rabello MM, Sterling FH et al. Diagnostic value of thyrotrophin-releasing hormone in pituitary and hypothalamic diseases. Ann Intern Med 1974;81:751–757.

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887 Woolf PD, Jacobs LS, Donofrio R. Secondary hypopituitarism: Evidence for continuing regulation of hormone release. J Clin Endocrinol Metab 1974;38: 71–76. 888 Nakasu Y, Nakasu S, Handa J, Takeuchi J. Amenorrhea-galactorrhea syndrome with craniopharyngioma. Surg Neurol 1980;13:154–156. 889 Leramo OB, Booth JD, Zinman B et al. Hyperprolactinemia, hypopituitarism and chiasmal compression due to carcinoma metastatic to the pituitary. Neurosurgery 1981;8:477–480. 890 Barbieri RL, Cooper DS, Daniels GH et al. Prolactin response to thyrotropin-releasing hormone (TRH) in patients with hypothalamic–pituitary disease. Fertil Steril 1985;43:66–73. 891 Bevan JS, Burke CW, Esiri MM, Adams CBT. Misinterpretation of prolactin levels leading to management errors is patients with sellar enlargement. Am J Med 1987;82:29–32. 892 Albuquerque FC, Hinton DR, Weiss MH. Excessively high prolactin level in a patient with a nonprolactin-secreting adenoma. J Neurosurg 1998;89:1043– 1046. 893 Tindal JS, Knaggs GS. Pathways in the forebrain of the rat concerned with the release of prolactin. Brain Research 1977;119:211–221. 894 Malven PV. Prolactin release induced by electrical stimulation of the hypothalamic preoptic area in unanesthetized sheep. Neuroendocrinology 1975;18:65–71. 895 Martin TL, Kim M, Malarkey WB. The natural history of idiopathic hyperprolactinemia. J Clin Endocrinol Metab 1985;60:855–858. 896 Sluijmer AV, Lappöhn RE. Clinical history and outcome of 59 patients with idiopathic hyperprolactinemia. Fertil Steril 1992;58:72–77. 897 Pontiroli AE, Falsetti L. Development of pituitary adenoma in women with hyperprolactinaemia: clinical, endocrine, and radiological characteristics. Brit Med J 1984;288:515–518. 898 Schlechte J, Dolan K, Sherman B et al. The natural history of untreated hyperprolactinemia: a prospective analysis. J Clin Endocrinol Metab 1989;68: 412–418. 899 Rjosk HK, Fahlbusch R, von Werder K. Spontaneous development of hyperprolactinaemia. Acta Endocrinol 1982;100:333–336. 900 Molitch ME, Reichlin S. Neuroendocrine studies of prolactin secretion in hyperprolactinemic states. In: Mena F, Valverde-R, C, eds. Prolactin Secretion: A Multidisciplinary Approach. New York: Academic Press, 1984:393–421. 901 Frantz AG. Endocrine diagnosis of prolactin-secreting pituitary tumors. In: Black PM, Zervas NT, Ridgway EC, Martin JB, eds. Secretory Tumors of the Pituitary Gland. New York: Raven Press, 1984:45–52. 902 Vance ML, Thorner MO. Prolactinomas. Endocrinol Metab Clinics N Amer 1987;16:731–753.

C h a p t e r

5 Thyroid-stimulating Hormone Virginia D. Sarapura Mary H. Samuels E. Chester Ridgway

TSH SUBUNIT GENES

INTRODUCTION Thyroid-stimulating hormone (TSH) is a glycoprotein produced by the thyrotrope cells of the anterior pituitary gland. TSH and two other pituitary hormones, luteinizing hormone (LH) and follicle-stimulating hormone (FSH), as well as the placental hormone chorionic gonadotropin (CG), consist of a heterodimer of two noncovalently linked subunits, a and b. The a-subunit is common to all four glycoproteins, whereas the b-subunit is unique to each and confers specificity of action. Each TSH subunit is encoded by a separate gene located on a different chromosome and is transcribed in a coordinated manner responsive mainly to the inhibitory effect of thyroid hormone. Production of bioactive TSH involves a process of cotranslational glycosylation and folding that enables combination between the nascent a- and b-subunits. TSH is stored in secretory granules and is released into the circulation in a regulated manner responsive mainly to the stimulatory effect of hypothalamic thyrotropin-releasing hormone (TRH). Circulating TSH binds to specific cell-surface receptors on the thyroid gland where it stimulates the production of thyroid hormones, L-thyroxine (T4) and L-triiodothyronine (T3), which act on multiple organs and tissues to modulate many metabolic processes as well as result in a feedback inhibition of TSH output. The introduction of sensitive TSH assays has allowed accurate measurement of the level of circulating TSH and has led to the recognition of abnormal production of TSH related with abnormal function of the thyroid gland reflecting in a wide spectrum of metabolic derangements.

The TSH b- and a-subunits are encoded by two separate genes located on different chromosomes. Since the first descriptions of the structures of these genes within the last decade, much information has been gained regarding the molecular events that result in the regulated production of TSH b- and a-subunit mRNAs. Most of this information has been obtained from studies performed in mouse thyrotropic tumors and rodent pituitary glands. Thyrotrope cells are believed to contain specific transcription factors that bind to the regulatory regions of the genes and interact with ubiquitous factors to initiate transcription. The identification of those specific factors is an active area of investigation. Regulation of TSH subunit gene transcription, mainly by thyroid hormone and TRH, is achieved by modulating the activity of the specific and ubiquitous factors.

Ontogeny of TSH Subunit Gene Expression TSH production is limited to the thyrotrope cells of the anterior pituitary gland. Studies performed in mice have shown that this restricted production is the result of a tightly controlled process of differentiation of pituitary cells beginning early in embryogenesis. Pluripotential cells differentiate into cells that produce specific types of pituitary hormones, including TSH, LH, FSH, adrenocorticotrophic hormone (ACTH), growth hormone (GH) and prolactin. The glycoprotein hormone a-subunit is the first pituitary hormone gene expressed [1]. This transcript is detected in the region of ectoderm that forms Rathke’s pouch, the origin of the pituitary gland, on mouse embryo day 11. The steps leading to the initiation of a-subunit expression are 172

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just beginning to be elucidated. Recent studies have shown that members of the Wnt and BMP gene families, specifically Wnt5a and BPM4, which are expressed in the adjacent neuroepithelium during the initial development of Rathke’s pouch, provide critical signals resulting in a-subunit expression throughout the developing pituitary gland [2]. In addition, the homeobox genes Rpx [3] and Ptx [4–6], as well as the LIM homeodomain factor P-LIM/Lhx3 [7–9], all of which are expressed in Rathke’s pouch before embryonic day 10, may play an activating role in the onset of a-subunit expression. The initiation of TSHb-subunit expression occurs in the rostral tip of the developing pituitary around mouse embryo day 12.5. This expression correlates temporally and spatially with thyrotrope embryonic factor (TEF), which is restricted to the pituitary at this embryonic stage, but in the mature organism it is expressed ubiquitously [10]. Expression in the rostral tip is found in the pituitary glands of both the wild type and the Snell dwarf mouse, a model of combined pituitary hormone deficiency lacking thyrotropes, lactotropes and somatotropes [11,12]. By birth, TSH b-subunit expression in the rostral tip has disappeared. However, another population of thyrotropes arises in the caudomedial region of the developing mouse anterior pituitary on embryonic day 15.5, one day after the expression of Pit-1, a pituitary-specific transcription factor, is detected in this region [13]. Pit-1 is a POU-homeodomain protein which was first described as an activator of the GH and prolactin genes [14]. Both Pit-1 and TSH b-subunit expression in the caudomedial region are present in the wild type but not in the Snell dwarf mouse, which has a mutation of the Pit-1 gene [15]. These data suggest that the second population of thyrotropes, associated with Pit-1 expression, may be the source of mature thyrotropes. Moreover, Pit-1 mutations have also been reported in humans [16–19] and are associated with a lack of thyrotropes, somatotropes and lactotropes, analogous to the Snell dwarf mouse phenotype. It has recently been shown that Pit-1 synergizes with PLIM/Lhx3, specifically the Lhx3a isoform, to activate the TSH b-subunit promoter [20]. Pit-1 expression, in turn, depends on the expression of another transcription factor, Prophet of Pit-1 (PROP-1). Mutations in this factor have been associated with many cases of combined pituitary hormone deficiency in humans, affecting not only thyrotrope, somatotrope and lactotrope but also gonadotrope expression [21] (see p. 197). Mutations in the PROP-1 gene were also found in the Ames dwarf mouse that exhibits a similar phenotype [22]. However, other factors must be involved in the initiation of thyrotrope-specific gene expression, since the presence of both Pit-1 and P-LIM/Lhx3 in somatotropes and lactotropes does not result in TSH production by these cells. Recent studies have suggested that another transcription factor, GATA-2, which has been found to synergize with Pit-1 to activate the TSH b-subunit promoter [23] (see p. 174) plays a critical role in thyrotrope differentiation [24].

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The hypothesis is that a ventral-dorsal gradient of GATA-2 occurs early in development in response to BMP-2: the intermediate cells that express both GATA-2 and Pit-1 activate the thyrotrope-specific genes, whereas the more ventral cells that express GATA-2 and not Pit-1 become gonadotropes, and the more dorsal cells that express Pit-1 and not GATA-2 become somatotropes and lactotropes (Fig. 5.1) [24].

TSH b-subunit Gene Structure The human TSH b-subunit gene has been isolated and its structure characterized [25–27]. This gene is 4.9 kilobase pairs (kb) in size, is located on chromosome 1 locus p22 [28,29] and is present as a single copy gene. The gene structure consists of three exons and two introns (Fig. 5.2). The first exon has 37 base pairs (bp) and contains the 5¢ untranslated region of the gene. It is separated from the second exon by a large first intron of 3.9 kb. The coding region of the gene is contained in the second (163 bp) and third (326 bp) exons, which are separated by a 0.45 kb intron, and the 3¢ untranslated region is contained in the third exon. This genomic organization is similar to that found in the mouse [30,31] and rat [32], with the exception that in the mouse TSH b-subunit gene the 5¢ untranslated region is contained in 3 exons resulting from additional splicing sites that are not present in the human TSH b-subunit gene [27,33]. Optional splicing of these exons in the mouse gene is a possible regulatory step that does not appear to be present in the human gene. In addition, the human exon 1 is 10 bp longer than that of the mouse and rat, presumably due to an insertion that displaces the TATA box, a sequence

Pit-1 GATA-2

S/L Tr T G

FIGURE 5.1. Thyrotrope cell origin during anterior pituitary development. Schematic representation of the regions of the pituitary where Pit-1 (hatched area) and GATA-2 (shaded area) transcripts are detected. Somatotropes and lactotropes (S/L) evolve from Pit-1(+)/GATA-2(-) cells, gonadotropes (G) from Pit-1(-)/GATA2(+) cells, and thyrotropes (T) from Pit-1(+)/GATA-2(+) cells [24]. The thyrotropes in the rostral tip (Tr) appear at an earlier stage in the region where TEF is expressed, but do not persist in the adult. From Dasen et al. [24].

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that is important for localizing RNA polymerase II activity, 9 bp further upstream relative to the TATA box in the mouse and rat genes.

TSH b-subunit Promoter As found in many other genes, sequences close to the transcriptional start site in the 5¢ flanking region of the TSH b-subunit gene contain elements responsible for initiating transcription and regulating the expression of the gene (Fig. 5.3). The transcriptional start site, defined by primer extension or S1 nuclease analysis, defines the start of the first exon [25–27]. A consensus TATA box is located 28 bp upstream of the transcriptional start site. In contrast to the human gene, both the mouse and the rat genes have two transcriptional start sites. The downstream transcriptional start site is homologous to that in the human gene and is responsible for most of the transcriptional activity (greater than 90%). The other transcriptional start site is located 40 bp

FIGURE 5.2. Schematic representation of the human thyroid-stimulating hormone (TSH) b-subunit gene. The bars indicate the three exons and the lines, which are not drawn to scale, represent intronic DNA. The solid areas represent the coding regions, the open areas represent the untranslated regions. The intronic DNA is removed by splicing to form mature mRNA. The numbers indicate the size in nucleotides of each exon or intron.

upstream and is also preceded by a TATA box. There is one bp difference between the rodent and human genes, that changes this upstream TATA box from TATATAA in the rodent to TGTATAA in the human, and it has been suggested that this difference may account for the lack of activity of the upstream transcriptional start site in the human gene [27]. Cell-specificity of TSH b-subunit Gene Expression

The lack of a reliable TSH-producing cell line has hindered investigation of the factors that determine thyrotropespecific expression. The tissues available for such studies have been limited to primary cultured cells from hypothyroid rodent pituitary glands and mouse thyrotropic tumors. Recently, the TaT-1 cell line was derived from a pituitary tumor produced by targeting T antigen expression in a transgenic mouse using the a-subunit promoter [34]. This cell line has been shown to express the TSH b-subunit and some of the other thyrotrope-specific genes, albeit at low levels [35]. More studies are needed to characterize this cell line. Using mouse thyrotropic tumors, studies have demonstrated that mouse TSH b-subunit gene region from -271 to -80 relative to the major transcriptional start site is sufficient to confer thyrotrope-specific activity [36] and that protein factors from thyrotropic tumors are able to bind to sequences within this region [37]. The pituitary transcription factor Pit-1 has been identified as one of the factors that binds to the human TSH bsubunit promoter between -128 and -58 [38] (Fig. 5.3). Pit-1 has also been found to bind to the mouse [39] and rat [40] TSH b-subunit promoters in regions important for activation. Haugen et al. had demonstrated that a thyrotrope-specific form of Pit-1, Pit-1T, is a strong transactivator of the mouse TSH b-subunit promoter [41,42], although Pit-1T is present in relatively small amounts and a human equivalent of Pit-1T has not been found. Recently, studies have shown that the transcription factor GATA-2 synergizes with Pit-1, resulting in a strong transactivation of the mouse TSH b-subunit promoter [23,43]. This synergy is mediated by partially overlapping sites located between -133 and -88.

FIGURE 5.3. Schematic representation of the human thyroid-stimulating hormone (TSH) b-subunit promoter. The transcriptional start site is indicated by the arrow and the TATA box is shown. The numbers above the line denote the position of the nucleotides relative to the transcriptional start site set at +1. The boxes under the line indicate the regions important for the responses to T3 (white boxes), Pit-1 (black boxes), thyrotropin-releasing hormone (TRH) (dotted boxes) and GATA-2 (crosshatched box). The GATA-2 binding sites have only been described in the mouse TSH b-subunit gene.

Chapter 5 Regulation of TSH b-subunit Gene Expression

Thyroid Hormone T3 inhibits TSH production primarily by its action at the transcriptional level (p. 178). T3 response elements (Fig. 5.3) are located between +3 and +37, in the first exon of the human TSH b-subunit gene [44]. There are two T3 receptor (TR) binding sites within this region, one from +3 to +13 and another from +28 to +37. Both sites are necessary for T3 inhibition of the human TSH bsubunit gene. The upstream site has a higher affinity for TR and is capable of binding TR dimers, whereas the downstream site binds only monomers with lower affinity. T3 response elements in the mouse [36] and rat [45,46] TSH b-subunit genes are also located near the transcriptional start site. The proximity of these response elements to the transcriptional start site suggests that T3 inhibition may be due to an interference with the activity of the transcriptional initiation complex. Bodenner et al. described the modulation of T3 inhibition by AP1, the dimerization product of the proto-oncogenes c-fos and c-jun [47]. These proto-oncogenes appear to interact with the sequence from -1 to +6 (TGGGTCA) that is homologous to the AP1 consensus binding site (TGAGTCA) (Fig. 5.4). Hypothalamic Factors TRH stimulates TSH production at various levels of synthesis and secretion, with a significant effect on the transcription of both TSH b- and a-subunits (see p. 179). Two TRH-response regions are located from -128 to -61 and from -28 to +8 of the human TSH bsubunit gene promoter [48] (Fig. 5.3). The upstream region contains binding sites for the pituitary-specific transcription factor Pit-1, suggesting a role for this or a similar factor in the regulation of the TSH b-subunit gene by TRH. In the rat TSH b-subunit gene, responsiveness to TRH has been localized to regions upstream of -204, where Pit-1 binding sites are also found [49,50]. Dopamine inhibits TSH b- and a-subunit gene transcription by decreasing the intracellular levels of cAMP, which acts as a stimulator (see p. 179). Studies of the TSH b-subunit gene have localized two regions of the promoter necessary for cAMP stimulation, from -128 to -61 bp and

FIGURE 5.4. Sequence comparison of the thyroidstimulating hormone (TSH) b-subunit region from -1 to +6 and consensus binding elements for thyroid hormone (T3), cAMP, phorbol esters (PhEst) and the c-jun/c-fos heterodimer (AP1). Spaces were introduced to maximize homology.

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from +3 to +8 bp [51]. The upstream region coincides with the TRH-responsive region and contains Pit-1 binding sites. The downstream region resides within the regions responsive to T3 (+3 to +37) and TRH (-28 to +8). The downstream region also overlaps with an AP1 binding site (-1 to +6). The sequence from -1 to +6 appears to cooperate with Pit-1 in mediating responses to cAMP and TRH [52]. Thus, multiple interactions between transcription factors and hormonal regulators appear to converge on the sequence from -1 to +6 (TGGGTCA), which is critically located close to the transcriptional start site, is similar to T3, cAMP, phorbol ester and AP1 response elements (Fig. 5.4), and is functionally important for the response to each of these regulators. Steroid Hormones Glucocorticoids inhibit TSH production but TSH subunit messenger RNA (mRNA) levels do not change significantly [53]. Their major effect may be at the secretory level (see p. 187). Testosterone has been shown to increase TSH b-subunit mRNA in castrate rat pituitary and mouse thyrotropic tumor [54]. Elements of the TSH b-subunit that respond to steroid hormones have not yet been described.

a-subunit Gene Structure The human glycoprotein hormone a-subunit gene is located on chromosome 6 [55] and is present as a single copy gene that is 13.5 kb in size and contains four exons and three introns [56] (Fig. 5.5). The first exon (94 bp) contains 5¢ untranslated sequences and is separated from the

FIGURE 5.5. Schematic representation of the human glycoprotein hormone a-subunit gene. The bars indicate the four exons and the lines, which are not drawn to scale, represent intronic DNA. The solid areas represent the coding regions, the open areas represent the untranslated regions. The intronic DNA is removed by splicing to form mature mRNA. The numbers indicate the size in nucleotides of each exon or intron.

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second exon by a 6.4 kilobase (kb) intron. The second exon contains 7 bp of 5¢ untranslated sequence and 88 bp of the coding region. The coding sequence continues in the third (185 bp) and fourth (75 bp) exons and the 3¢ untranslated region (220 bp) is contained in the fourth exon. The second and third introns are 1.7 kb and 0.4 kb, respectively. The genomic organization of the mouse [57], rat [58] and cow [59] a-subunit genes are similar, except that in the rat and cow the second intron is located 12 bp downstream, resulting in a peptide sequence that is 4 amino acids longer. There are also differences in the length of the 5¢ untranslated sequence, which is 10 bp longer in the mouse, apparently due to a 10 bp insertion between the TATA box and the transcriptional start site.

a-subunit Promoter The elements responsible for initiating transcription and regulating the expression are located in the 5¢ flanking region of the human a-subunit gene (Fig. 5.6). The human a-subunit gene contains a concensus TATA box located 26 bp upstream of the transcriptional start site [56]. A single transcriptional start site has been found in the glycoprotein hormone a-subunit genes of all the species that have been studied. Cell-specificity of a-subunit Gene Expression

The glycoprotein hormone a subunit gene is unique in that it is expressed in thyrotropes, gonadotropes and placental cells, and in each of these cell types it is differentially regulated. Studies from several laboratories using the human and mouse genes have demonstrated that the cell-specific expression in each cell type is dependent on vastly different regions of the promoter (Fig. 5.6). Whereas the region downstream of -200 is sufficient for placental expression [60–62], gonadotropes require sequences between -225 and -200 [63], and regions further upstream appear to be critical for thyrotrope expression [51,64–67]. The elements involved in human placental a-subunit expression extend from -177 to -84 and include the upstream regulatory element (URE) also called trophoblast-specific element (TSE) that binds the placental-specific protein TSEB

[60,62], two cAMP response elements (CREs) [60,61,68] that bind the ubiquitous protein CREB [69], the junctional regulatory element ( JRE) that binds a 50-kDa protein [70], the CCAAT-box that binds a 53-kDa a-subunit binding factor (a CBF) [71] and a GATA motif that interacts with GATA-binding proteins [72]. Some of these regions binding to similar factors may also play a role in pituitary a-subunit expression. A region from -225 to -200 that binds the orphan nuclear receptor SF-1 appears to be critical for gonadotrope expression of the a-subunit gene [63,73], but this region has no effect on thyrotrope expression [74]. Basic-helix-loop-helix E-box-binding proteins [75] and GATA-binding proteins [72] also appear to play a role in asubunit expression in gonadotropes. Transgenic mouse studies have shown that 313 bp of the bovine a-subunit 5¢ flanking DNA, which contain the SF-1-binding region, targeted expression to gonadotropes but not thyrotropes [76], suggesting that this region was sufficient for expression in gonadotropes. It was then shown that 480 bp of the mouse a-subunit 5¢ flanking DNA was able to target transgenic expression to both gonadotropes and thyrotropes [77], in agreement with in vitro transfection studies which showed that the same promoter region mediates a high level of expression in thyrotrope and gonadotrope cells [65,67]. Several sequences within the region from -480 to -300 appear to be important for mouse a-subunit expression in thryotropes but not gonadotropes [67]. Among these is the sequence from -434 to -421 that interacts with the developmental homeodomain transcription factor Msx1 [78]. This factor was found to be expressed in mature thyrotropes, but its role in a-subunit expression has not been elucidated. Another important sequence is the pituitary glycoprotein hormone basal element, or PGBE, extending from -342 to -329, that is critical for both thyrotrope and gonadotrope expression [7]. The PGBE interacts with P-LIM (also known as mLIM3 or Lhx3), a pituitary-specific LIMhomeodomain transcription factor [8], that is important not only for thyrotrope and gonadotrope cell-specification but is also important for somatotropes and lactotropes [9]. Other sequences within the 480 bp promoter have been found to interact with the pituitary-specific homeodomain factor Ptx-1 [5,67], and a synergism between Ptx-1 and P-LIM,

FIGURE 5.6. Schematic representation of the human glycoprotein hormone a-subunit promoter. The transcriptional start site is indicated by the arrow and the TATA box is shown. The numbers above the line denote the position of the nucleotides relative to the transcriptional start site set at +1. The boxes under the line indicate the regions important for the responses to T3 (TRE) and cAMP (CRE), and the placental-specific, gonadotrope-specific and thyrotrope-specific activities, as shown. The thyrotrope-specific regions other than the P-LIM binding sites have only been described in the mouse a-subunit gene.

Chapter 5

mediated by the coactivator C-LIM, has recently been described [79]. Notwithstanding the findings with the mouse a-subunit gene, other studies showed that 1.6 kb of 5¢ flanking DNA of the human a promoter expressed in gonadotropes but not in thyrotropes of transgenic mice [80], suggesting that either the mouse thyrotropic factors are unable to activate the human promoter, or else the sequences required for thyrotrope expression of the human a-subunit gene may be located further upstream than those of the mouse gene. Recent studies with the mouse promoter also showed that regions located approximately 4 kb upstream further enhanced transgenic expression in both thyrotropes and gonadotropes, by interacting with proximal sequences [74,77,81]. The active region was localized to 125 nucleotides upstream of -3700, and this same region was shown to mediate inhibition of expression in GH3 somatotropic cells [82]. In spite of significant advances in this area, thyrotrope-specific factors that determine a-subunit gene expression have not yet been described. In contrast to the TSH b-subunit gene, it has been shown that Pit-1 is not necessary for a-subunit gene expression [39]. Regulation of a-subunit Gene Expression

Thyroid Hormone T3 inhibition of a-subunit gene transcription is observed in thryotropes in coordination with that of TSH b-subunit (see p. 178). The T3 response element of the human glycoprotein hormone a-subunit gene promoter is located from -22 to -7 [83] (Fig. 5.6). Similar to the TSH b-subunit gene, the T3 response elements of the human as well as the mouse [51] and rat [58] a-subunit genes are located close to the transcriptional start site and the mechanism of inhibition is likely to involve interference with the activity of the transcriptional initiation complex. T3 inhibition may be mediated by different isoforms of the T3 receptor (TR) [84,85] in combination with the corepressors SMRT and NCoR [86]. Studies have suggested that mutations of the T3 response element of the human a-subunit promoter that eliminate TR binding do not abrogate the inhibitor effect of T3, suggesting that protein–protein interactions may be more important than protein-DNA binding [87]. Hypothalamic Factors TRH stimulates TSH production by effects at the transcriptional, posttranscriptional and secretory levels, discussed in later sections (see pp. 179 and 185). Although TRH has clearly been shown to stimulate a-subunit transcription, the TRH-responsive element of the human glycoprotein hormone a-subunit gene promoter has not yet been localized. Studies using primary cultures of rat pituitaries have shown that the human a-subunit promoter containing 846 bp of 5¢ flanking DNA responds to TRH with a 23% increase of activity [88], and that this stimulation is mediated by a region located just upstream of -200 [89]. However, this increase was not significant, at least in part due to the presence of a-subunit expressing gonadotropes in the mixed population of pituitary cells.

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Subsequent studies showed that a CRE-binding protein that binds to the region from -151 to -135 of the human asubunit promoter is important for TRH regulation, as well as a Pit-1-like protein that binds to a more distal region from -223 to -190 [90]. a-subunit gene expression in thyrotropes is inhibited by dopamine in coordination with the expression of the TSH b-subunit gene. Its action is mediated by decreases in intracellular cAMP levels. The CRE of the human glycoprotein hormone a-subunit gene promoter extends from -146 to -111 and has been well characterized by several investigators [61,68] (Fig. 5.6). The element consists of an 18 bp repeat containing a core palindrome TGACGTCA. A single bp change in this palindrome in the mouse, rat and cow genes to TGATGTCA significantly reduces cAMP responsiveness. The importance of the CRE in placental a-subunit gene expression has been established, and it is of interest in this respect that rodents and cows do not express a-subunit in the placenta. The finding that the equine a-subunit gene does express in the placenta but also lacks a competent CRE has led to the description of a GATA element located upstream of the CRE that confers both placental-specific expression and cAMP responsiveness through its interaction with a-ACT, a placental protein related to the GATAbinding factors [91]. In gonadotropes, the interaction of the CRE with the transcription factors ATF2 and c-Jun appears to be functionally important [92], as well as the GATAbinding proteins [72]. The importance of those same interactions in thyrotropes has not been established. Steroid Hormones Regulation of TSH transcription by steroid hormones is probably of limited importance. Several studies have described the steroid hormone regulation of asubunit gene transcription and have found that in general the effects are inhibitory and correlate more closely with physiologic effects seen in gonadotropes. Androgen inhibition and androgen receptor (AR) binding has been localized to a region from -120 to -100 that coincides with the JRE described above [93]. Negative regulation by estrogen was described in the gonadotropes of transgenic mice expressing the cholamphenicol acetyl-transferase gene under the control of the human (1.5 kb) and bovine (314 bp) 5¢ flanking regions, but no binding of these regions to the estrogen receptor (ER) was detected, suggesting that estrogen acts indirectly [94]. This was also suggested by transient transfection studies that failed to detect an effect of estrogen on the human a-subunit gene promoter activity in gonadotrope cells [93]. However, other studies using rat pituitary cells of somatomammotrope origin have found positive regulation by estrogen localized to the proximal 98 bp of 5¢ flanking DNA of the human a-subunit gene and binding of the ER to the T3 response element from -22 to -7 [95]. Transcriptional inhibition by glucocorticoids appears to be mediated by binding of the glucocorticoid receptor (GR) to sequences between -122 and -93 of the human a-subunit gene [96]. This region contains the JRE

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and binds to the AR, as described above. However, no direct binding was detected in other studies, suggesting that the GR inhibits transcription by interfering with other transactivating proteins [97]. BIOSYNTHESIS OF TSH The intact TSH molecule is a heterodimeric glycoprotein with a total molecular weight of 28-kDa that is composed of the noncovalently linked a- and b-subunits. TSH biosynthesis and secretion by thyrotropes of the anterior pituitary are finely regulated events. This section examines our understanding of the biosynthesis of TSH, including the processes of transcription, translation, glycosylation, folding, combination and storage. Excellent reviews on these subjects have been published [98–100].

Transcription of TSH The TSH b- and a-subunit genes are transcribed as directed by their respective promoters in the presence of both ubiquitous and specific transcription factors as described above. The transcribed RNAs undergo a process of splicing at the exon–intron junctions that leads to the production of the mature mRNAs that then exit the nucleus and are translated in the cytoplasm. Transcription of the TSH b- and a-subunit genes is coordinated under the influence of physiologic regulators, the most important of which are T3 and TRH. Thyroid Hormone Regulation

TSH b- and a-subunit gene transcription rates are markedly inhibited by T3. Studies using mouse thyrotropic tumors have demonstrated that suppression of TSH b- and asubunit mRNA transcription rates measured by nuclear run-on assays is evident by 30 minutes after treatment and is maximal by 4 hours [101] (Fig. 5.7, upper panel). This effect was seen in the presence of the protein synthesis inhibitor cycloheximide, indicating that it did not require the induction of an intermediary protein [102] (Fig. 5.7, lower panel). Other studies using mouse and rat pituitaries and mouse thyrotropic tumors have demonstrated that steadystate mRNA levels of TSH b- and a-subunit are dramatically decreased by T3 [103–106]. In the hypothyroid state, the ratio of a- to TSH b-subunit mRNA ranges from 0.35 to 1.28. In the euthyroid state, both subunits decreased, with the ratio changing from 1.1 to 1.7. In the hyperthyroid animal, inhibition by T3 was greater for the TSH b-subunit mRNA, that decreased to undetectable levels, whereas asubunit mRNA decreased to 20% to 25% of control so that the ratio of a- to TSH b-subunit mRNA increased to approximately 2–3. Some studies in humans and animals have shown that TSH paradoxically increased after T3 treatment [107–110], although other studies did not show it [104,105,111,112]. When this effect was observed, it occurred early in the treatment of a severe, prolonged

FIGURE 5.7. T3 inhibition of thyroid-stimulating hormone (TSH) subunit transcription. Upper panel, the time course of TSH b(shaded bars) and a-subunit (white bars) mRNA synthesis. Mice bearing thyrotropic tumors (2 per group) were treated for up to 5 days with daily T3 injections. Tumors were removed after the times indicated and the transcription rates were measured in isolated nuclei. From Shupnik et al. [101]. Lower panel, the effect of cycloheximide on T3 inhibition of TSH b- and a-subunit mRNA synthesis. Mouse thyrotropic tumor explants were placed in culture for up to 4 hours and treated with 5 nM T3 in the presence or absence of the protein synthesis inhibitor cycloheximide, then transcription rates were measured in isolated nuclei. The values represent the mean ± SEM of four determinations performed in duplicate. From Shupnik et al. [102].

hypothyroid state and may be due to defective synthesis in this state that is corrected by low levels of T3. The mechanism of action of T3 involves interaction with nuclear receptors that act mainly at the transcriptional level. The transcriptional response to T3 is proportional to the nuclear receptor occupancy [112], and the time course of T3 nuclear binding and transcriptional inhibition are also in agreement [102] (Fig. 5.8). It is now believed that TR are bound to the DNA response elements in the absence of ligand, and that T3 binding appears to modify the interaction between the TR and the response element as well as

Chapter 5

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FIGURE 5.8. The effect of thyroid hormone on the transcription of the thyroidstimulating hormone (TSH) b- (black circles) and a-subunit (white circles) genes. Murine thyrotropic tumor explants were incubated for up to 4 hours with 5 nmol T3 for transcription measurements or with 5 nmol [125I]T3 with or without 1,000-fold excess of unlabeled T3 for binding measurements. Transcription rates were measured in isolated nuclei. There is an inverse relationship between T3 binding and TSH b- and a-subunit mRNA synthesis. From Shupnik et al. [102].

with other transcription factors [113]. It has been shown in mouse and rat thyrotropes that there are several types of TR: TRa1, TRb1 and TRb2, as well as non-T3 binding variant a2 [114,115]. These isoforms are able to bind DNA as monomers, homodimers and heterodimers of two different TR isoforms. Recent studies demonstrated that TRb2, which is expressed predominantly in the pituitary and T3responsive TRH neurons [116], appears to be the most important for the regulation of TSH [117,118]. Moreover, TRb2 deficient mice had a phenotype consistent with pituitary resistance to thyroid hormone, with increased TSH and thyroid hormone levels, even in the presence of TRb1 and TRa1 [119,120]. However, TRb1 and TRa1 may still play a role, since they are able to form heterodimers with TRb2. Heterodimers of a TR and a TR accessory protein, such as 9-cis retinoic acid receptors (RXR), may also bind to DNA [121], constituting heterodimeric complexes that may have different affinities for specific DNA sequences and different functional actitivies. A particular RXR isoform, RXRg1, is uniquely expressed in thyrotropes and appears to mediate the inhibition by 9-cis-retinoic acid through a region extending from -200 to -149 of the mouse TSH b-subunit promoter [122]. Other proteins that interact with TR include the coactivators, such as the glucocorticoid receptor interacting protein-1 (GRIP-1) and the steroid receptor coactivator-1 (SRC-1) [123], and corepressors, such as the silencing mediator for retinoid receptors and thyroid hormone receptors (SMRT) and the nuclear receptor corepressor (NCoR) [86,124]. These coactivators and corepressors modulate the effect of many members of the steroid–thyroid hormone receptor superfamily. Their role in the regulation of the TSHb-subunit promoter by thyroid hormone remains to be elucidated.

Posttranscriptional effects of T3 have also been described. T3 decreases the half-life of TSH b-subunit mRNA and decreases the size of the poly(A) tail [125]. The shortening of the poly(A) tail is thought to cause mRNA instability. This effect of T3 was observed for TSH b- but not for a-subunit mRNA, but this may have been due to the fact that the population of gonadotropes present in the pituitary sample may have masked the effect on thyrotropic a-subunit mRNA. Leedman et al. showed that T3 increased the binding of an RNA-binding protein present in rat pituitary to the 3¢ untranslated region of the rat TSH b mRNA [126] and also induced a shortening of the poly(A) tail of the mouse TSH b mRNA from 160 to 30 nucleotides [127]. Hypothalamic Regulation

TRH A direct effect of TRH on the transcription rates of both TSH b- and a-subunit genes has been demonstrated in primary cultures of hypothyroid rat pituitary cells [128]. The maximal stimulation ranged from two- to fivefold and occurred after 30 minutes of treatment with 10-7 M TRH for both the TSH b- and a-subunit genes (Fig. 5.9). TRH acts on a cell-surface receptor [129,130]. Studies in rat pituitary somatomammotrope cells, where TRH stimulates prolactin production, have suggested that phosphatidylinositol, protein kinase C and calcium-dependent pathways may be involved [131–133] (see TSH Secretion), and as described in a previous section, TRH stimulation of the TSH b-subunit promoter may be mediated by AP1 [52] (see pp. 175 and 177). Dopamine Dopamine acts by decreasing the levels of intracellular cAMP. This effect is achieved by binding to cell-surface receptors [134] that inhibit adenylate cyclase

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Translation of TSH

FIGURE 5.9. The effect of thyrotropin-releasing hormone (TRH) on thyroid-stimulating hormone (TSH) subunit mRNA synthesis. Pituitary cells (3–5 ¥ 106 cells/60 mm plate) were treated with 100 nM TRH for up to 4 hours, then transcription rates were measured in isolated nuclei. The values represent the mean ± SEM of four determinations. From Shupnik et al. [128].

activity [135,136]. The stimulatory effect of cAMP on TSH b- and a-subunit gene transcription was demonstrated indirectly by reversal with cAMP of the 65–70% inhibition of TSH b- and a-subunit mRNA synthesis caused by dopamine [128] and directly by treatment of primary pituitary cultures with forskolin, an activator of adenylate cyclase [137]. Somatostatin (SRIH) The inhibitory effect of SRIH on TSH production occurs mainly at the secretory level (see p. 186). Studies in GH3 somatomammotrope cells have shown that SRIH inhibits the cAMP/protein kinase A pathway [138], resulting in an inhibition of transcription. However, a direct effect on TSH gene transcription has not been demonstrated. Steroid Hormone Regulation

The impact of glucocorticoids on TSH production occurs mainly at the secretory level (see p. 188) while effects on TSH subunit gene transcription have not been observed [53,139]. Androgens have been shown to increase TSH bsubunit mRNA levels in the pituitary of castrate male rats [54] and decrease these levels in intact female rats [140]. Estrogens, on the other hand, do not appear to affect TSH transcription directly, but they increase the number of pituitary nuclear receptors for T3, and augment T3 suppression of TSH subunit gene transcription [141,142]. Estrogens may also indirectly affect TSH production, by increasing the expression and stability of TRH receptors [143], but this has only been demonstrated in GH3 somatotrope cells, and has not been described in thyrotrope cells.

The next series of processes of TSH biosynthesis are summarized in Fig. 5.10 [144]. In the cytoplasm, the mRNAs for TSH b- and a-subunit are independently translated by ribosomes. The first peptide sequence produced consists of a “signal” peptide of 20 amino acids for TSH b and 24 amino acids for a [145]. This signal peptide is important for interaction of the ribosome with the rough endoplasmic reticulum and its hydrophobic nature allows insertion through the lipid bilayer of the membrane. Translation into TSH b and a presubunits continues into the lumen, where cleavage of the signal peptide occurs before translation is completed, resulting in the formation of a 118 amino acid TSH b-subunit [146] and a 92 amino acid a-subunit [56]. Cleavage of TSH b to a protein of 112 amino acids appears to be an artifact of purification. Synthesis of recombinant TSH b-subunit has resulted in two products of 112 and 118 amino acids, both of which are similarly active in vitro [147]. Mutations of the TSH b-subunit have recently been described in humans with the diagnosis of central hypothyroidism (see p. 197). In one kindred, translation of TSH is affected by a mutation that introduces a premature termination codon [148]. This results in a truncated peptide of only 11 amino acids that is biologically inactive.

Glycosylation of TSH Glycosylation of TSH is not essential for receptor binding but has a significant impact on its biological activity [149,150]. The TSH b-subunit has a single glycosylation site, the asparagine residue at position 23, whereas the a-subunit is glycosylated in two sites, the asparagine residues at positions 52 and 78 [151] (Fig. 5.11). Excess free a-subunit is glycosylated at an additional site, the threonine residue at position 39 [152]. This residue is located in a region believed to be important for combination with the TSH b-subunit. It is not known whether glycosylation at this residue is a regulated step that inhibits combination with the TSH b-subunit or it occurs in excess free a-subunits because this site is exposed. Extensive studies on the processes of TSH subunit glycosylation have been carried out. Glycosylation of the TSH b- and a-subunits begins before translation is completed (cotranslational glycosylation), while addition of the second oligosaccharide in the a-subunit occurs after translation is completed (posttranslational glycosylation). The first step in this process involves the assembly of a 14-residue oligosaccharide, (glucose)3-(mannose)9-(N-acetylglucosamine)2 on a dolichol-phosphate carrier. This oligosaccharide is then transferred to asparagine residues by the enzyme olygosaccharyl transferase that recognizes the tripeptide sequence (asparagine)-(X)-(serine or threonine) [153]. This mannoserich oligosaccharide is progressively cleaved in the rough endoplasmic reticulum and Golgi apparatus. An intermedi-

; Chapter 5

RER

Proximal golgi

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α

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α

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α

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H2N H2N

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Distal golgi

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FIGURE 5.10 . Biosynthesis of thyroid-stimulating hormone (TSH). Schematic representation of the processes of translation and glycosylation within the rough endoplasmic reticulum (RER) and golgi apparatus, divided into proximal and distal. Cleavage of the aminoterminal (H2N) signal peptide and early addition of high mannose chains (black boxes) as well as combination of a- and b-subunits occur in the RER. In the proximal golgi, oligosaccharide chains are modified (shaded boxes) and the final steps of sulfation and sialation occur in the distal golgi apparatus. From Weintraub and Gesundheit [144].

FIGURE 5.11. Oligosaccharide chains of thyroid-stimulating hormone (TSH). Schematic representation of the typical oligosaccharide chains present on the TSH heterodimer and the free a-subunit. Glycosylated asparagine (Asp) and threonine (Thr) residues are indicated. Black and white circles, half-circles and triangles represent the oligosaccharide chain residues as indicated in the key. From Weintraub and Gesundheit [144].

ate with only six residues is produced, and then other residues are added resulting in complex oligosaccharides [154]. The residues added include N-acetylglucosamine, fucose, galactose and N-acetylgalactosamine. Oligosaccharides prior to the six-residue intermediate are termed highmannose and are sensitive to endoglycosidase H, that releases the oligosaccharide from the protein, whereas the intermediate and the complex oligosaccharides are endoglycosidase H-resistant. Complex oligosaccharides usually consist of two branches (biantennary) but sometimes three or four branches are seen, as well as hybrid oligosaccharides consisting of one complex and another high-mannose branch. Sulfation and sialation occur late in the pathway, within the distal Golgi apparatus [155]. Sulfate is bound to N-acetylgalactosamine residues, and sialic acid, or its precursor N-acetylneuraminic acid, is bound to galactoside residues

[99]. Thus, the activation of the enzymes sulfotransferase and N-acetylgalactosamine transferase may be important regulatory steps that impact the ratio of sulfate to sialic acid. As demonstrated with LH, it appears that sulfation increases and sialylation decreases the bioactivity of TSH [99], since the exclusively sialylated recombinant glycoprotein produced in Chinese hamster ovary cells has been found to have attenuated activity in vitro [156,157]. Processing of complex oligosaccharides appears to occur at a slower rate for secreted glycoproteins, such as TSH, when compared to non-secreted glycoproteins. For example, after an 11-minute pulse labeling with [35S]methionine and a 30-minute chase only a few a-subunits were endoglycosidase H resistant and only 76% reached this stage after an 18-hour chase [158]. Secretion was observed after a 60-minute chase and the secreted products—TSH, free

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a-subunit but no free b-subunit—had mostly complex oligosaccharides associated to them [151]. It may be important to note that many of the studies described were carried out in thyrotropic tumor tissue obtained from hypothyroid mice, and glycosylation may differ in the euthyroid as compared to the hypothyroid state. In addition, differences between species have been noted, such as the human TSH containing more sialic acid than the bovine TSH [146].

αL3

αL1

α64-81

α11-20

Regulation of TSH Glycosylation

Glycosylation is a regulated process that is primarily modulated by TRH and thyroid hormone [159]. Primary hypothyroidism [160,161] and TRH administration [162,163] have been found to increase oligosaccharide addition that results in an increased bioactivity of TSH [164] (see p. 190). The same was noted in patients with resistance to thyroid hormone [165]. TSH glycosylation patterns were also found to differ in several pathological states, such as central hypothyroidism, TSH-producing pituitary adenomas and euthyroid sick syndrome [166,167]. Also observed were changes in the sulfation and sialylation of the oligosaccharide residues, which modulates bioactivity [155,161,168,169] (see p. 190).

α88-92

α33-38

β88-95

α52

α40-46

Folding, Combination and Storage of TSH

The recent elucidation of the crystal structure of human CG (hCG) [170] has allowed the construction of a model of human TSH (Fig. 5.12), supported by other evidence [171,172]. This model has greatly facilitated the interpretation of structure–function studies of the protein backbone. However, crystallization was only achieved with partly deglycosylated hCG, so it is likely that the conformation of the glycosylated protein may differ to some extent, although nuclear magnetic resonance studies suggest that the asubunit carbohydrate moieties project outward and may be freely mobile [173]. Nevertheless, this model predicts that the tertiary structure of each TSH subunit consists of two hairpin loops on one side of a central knot formed by three disulfide bonds and a long loop on the other side. In this tertiary structure, the glycoprotein hormones share features in common with transforming growth factor b, nerve growth factor, platelet-derived growth factor, vascular endothelial growth factor, inhibin and activin, all of which are now grouped in the family of cystine knot growth factors [174]. Folding of nascent peptides begins before translation is completed. It has been shown that proper folding is dependent on glycosylation, since the drug tunicamycin that prevents the initial oligosaccharide transfer to the asparagine residue results in a peptide that does not fold properly and is degraded intracellularly [175]. Site-directed mutagenesis of a single glycosylation site also disrupted processing and decreased TSH secretion in transfected Chinese hamster ovary cells [176]. Folding is a critical step that allows correct internal disulfide bonding that stabilizes

β96-105

βL1

β58-69

βL3

FIGURE 5.12. Human thyroid-stimulating hormone (TSH) ribbon homology model showing domains important for activity. The schematic drawing is based on a molecular homology model [171] built on the template of the human chorionic gonadotropin (hCG) model derived from crystallographic coordinates obtained from the Brookhave Data Bank [170]. The a-subunit is shown as a checkered line, and the TSH b-subunit as a solid line. The two hairpin loops (L1, L3) in each subunit are marked. The long loops (L2) in each subunit extend from the opposite side of the central cystine knot. The functionally important a-subunit domains are boxed: a11–20, a33–38, a40–46 (“a helix”), a52, a64–81 and a88–92. The functionally important b-subunit domains are indicated within the line drawing: b58–69 (crossed line), b88–95 – the “determinant loop” or N-terminal segment of the seat-belt – (beaded line), and b96–105 – C-terminal segment of the seat-belt – (dashed line). The b-subunit beyond 106 is not drawn because the corresponding region of hCG was not traceable. The oligosaccharide chains are not shown because hCG was deglycosylated before crystallization. From Grossmann et al. [100].

the tertiary structure of the protein allowing subunit combination. Combination of TSH b- and a-subunits begins soon after translation is completed in the rough endoplasmic reticulum, and continues in the Golgi apparatus [151,177]. Subunit combination then accelerates and modifies oligosaccharide processing of the a-subunit [178]. In fact, studies

Chapter 5

have suggested that the conformation of the a-subunit differs after combination with each type of b-subunit [179], and this may affect subsequent processing. The rate of combination of TSH b- and a-subunits has been examined in mouse thyrotropic tumors. After a 20-minute pulse labeling with [35S]methionine, 19% of TSH b-subunits were combined with a-subunits, and this percentage increased to 61% after an additional 60-minute chase incubation [151,177]. The sequence of the TSH b-subunit from amino acid 27 to 31 (CAGYC) is highly conserved among species and is thought to be important for combination with the asubunit. In a case of congenital hypothyroidism, a point mutation in the CAGYC region (see p. 197) results in the synthesis of altered TSH b-subunits that are unable to associate with a-subunits, with consequent lack of intact TSH production [180]. A lack of free circulating TSH b-subunit was also observed, suggesting that combination with asubunit is necessary for TSH b-subunit secretion. This phenomenon was also demonstrated in studies where synthesis of wild-type recombinant TSH b-subunit was carried out in the presence or absence of recombinant a-subunit [181]. Using site-directed mutagenesis, another study showed that a mutation at residue 25 in the glycosylation recognition site that substitutes a serine for a threonine does not alter glycosylation but decreases TSH production by 70%, possibly because of disruption of the nearby CAGYC region [182]. After TSH and free a-subunit are processed in the distal Golgi apparatus they are transported into secretory granules or vesicles [183]. The secretory granules constitute a regulated secretory pathway, mainly influenced by TRH and other hypothalamic factors. These granules contain mostly TSH, whereas free a-subunit is contained in the secretory vesicles that constitute a nonregulated secretory pathway. TSH SECRETION TSH is synthesized and secreted only by thyrotrope cells in the anterior pituitary gland. Thyrotropes comprise approximately 5% of anterior pituitary cells, and are located chiefly in the anteromedial portion of the pituitary gland [184]. In healthy human subjects, the production rate of TSH is 100–400 mU/day [185–188]. The distribution space of TSH is slightly greater than the plasma volume, and the plasma clearance rate of TSH is approximately 50 ml/min [188]. In euthyroid subjects, the half-life of TSH in plasma is approximately 50 minutes [187]. In hypothyroid subjects, secretion rates of TSH increase up to 10 to 15 times normal rates, while the metabolic clearance rate decreases slightly. In contrast, in hyperthyroid subjects, secretion of TSH is suppressed, and metabolic clearance of exogenous TSH is accelerated [188]. The oligosaccharide chains of TSH appear to play important roles in TSH assembly, secretion, and action [144]. Nonglycosylated TSH subunits are degraded intracellularly before secretion, and inhibition of oligosaccharide chain

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processing interferes with intracellular TSH transport (see p. 182). The final structure of secreted TSH oligosaccharide is important in determining the biological activity and metabolic clearance rate of the circulating hormone. After deglycosylation, TSH has markedly reduced bioactivity and a more rapid metabolic clearance rate than the native hormone [189]. Variations in the oligosaccharide chains of TSH have been described in many clinical situations, and may affect biological half-life and bioactivity. For example, subjects with primary hypothyroidism secrete highly sialylated TSH isoforms with impaired bioactivity, and Lthyroxine treatment results in improvement in these properties [190].

Ontogeny of TSH Levels Immunoreactive TSH cells have been observed in the human pituitary gland by 12 weeks of gestation [191]. At approximately the same time, TSH is first detected in the pituitary and the serum [192–196]. TRH is measurable in hypothalamic tissue at 8 to 10 weeks, and the levels increase progressively to term. Serum and pituitary TSH levels remain low until week 18. At that point,TSH levels increase dramatically, followed by increases in serum T4 and T3 concentrations [192]. It appears that this increase in TSH represents the maturation of the hypothalamic–pituitary unit and the development of TRH input to the pituitary gland. There are continued increases in fetal serum TSH and T4 concentrations between 20 and 40 weeks of gestation. Pituitary TSH responsiveness to exogenous TRH occurs early in the third trimester, while negative feedback control of TSH secretion develops during the last half of gestation and the first 1 to 2 months of life [197]. Within 30 minutes of birth in term infants, there is an abrupt rise in serum TSH levels, followed by a marked increase in T3 concentrations within 4 hours and a lesser increase in T4 levels within the first 24 to 36 hours of extrauterine life. The initial increase in serum TSH levels appears to be stimulated by cooling the extrauterine environment. Serum TSH levels fall to the normal adult range by 3 to 5 days after birth, and serum thyroid hormone levels stabilize by 1 to 2 months of postnatal life. Healthy premature infants (less than 37 weeks gestational age) tend to have lower serum TSH levels at birth compared to term infants, although there is wide variability. There is a slight decrease in TSH levels in these infants during the first week of life, followed by a gradual increase to normal term levels. Serum TSH levels are lower at birth in ill premature infants compared to healthy premature infants matched for gestational age, but rise towards normal levels during recovery [198,199].

Patterns of TSH Secretion In healthy subjects, TSH is secreted from the pituitary gland in a dual fashion, with secretory bursts (pulses) superim-

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posed upon tonic (apulsatile) secretion (Fig. 5.13, upper panel). Tonic TSH secretion accounts for 30% to 40% of the total amount of TSH released into the circulation, while secretory bursts account for the remaining 60% to 70%. TSH pusles occur on average every 2 to 3 hours, although there is considerable variability among individuals, as well as variability due to blood sampling frequency and pulse analysis techniques [200,201]. Under normal conditions, TSH is secreted in a distinct circadian pattern, with nocturnal levels increasing up to twice daytime levels [197] (Fig. 5.13, upper panel). Peak TSH levels may occur at any time between 2300 and 500 hours, depending on the individual. In some healthy subjects, nocturnal TSH levels increase to slightly above the reported normal range for TSH assays. Lowest TSH levels occur at about 1100 hours. The TSH circadian rhythm is not apparent in premature or fullterm infants less than 4 weeks old, but begins to emerge between 1 and 2 months of life, and is well established in healthy children [202,203]. The circadian variation in TSH levels is due mainly to increased nocturnal TSH pulse amplitude, with some contribution from a slight increase in nocturnal TSH pulse frequency [201]. The nocturnal increase in TSH levels can precede the onset of sleep, and sleep deprivation actually enhances TSH secretion [200,204–206]. Therefore, in contrast to other pituitary hormones with a circadian variation, such as GH, the nocturnal rise in TSH levels is not sleep entrained. Instead, there appears to be a sleep-related inhibition of TSH release that is not of sufficient magnitude to counteract the stimuli that lead to the nocturnal TSH surge. Subjects with mild primary hypothyroidism continue to show a nocturnal rise in TSH pulse amplitude, while patients with severe primary hypothyroidism have markedly increased TSH pulse amplitude throughout the day, with no nocturnal increase (Fig. 5.13, middle panel) [201,207]. The normal TSH circadian variation is reestablished in these subjects by thyroxine replacement therapy [207]. In contrast, patients with hypothalamic–pituitary diseases and central hypothyroidism continue to secrete TSH in pulses, but lose the nocturnal TSH surge in pulse amplitude, and therefore secrete less TSH over a 24-hour period [208] (Fig. 5.13, lower panel). A similar pattern of reduced 24-hour TSH secretion due to lack of the normal nocturnal TSH surge is seen in critical illness, and may contribute to the low circulating thyroid hormone levels in severely ill patients [209]. To date, the origin of pulsatile and circadian TSH secretion is unknown. Thyroid hormones alter TSH pulse amplitude, but have little effect on pulse frequency [207], and therefore are unlikely to participate in TSH pulse generation. Some investigators have hypothesized that a TSH pulse generator exists in the hypothalamus, consisting of TRH neurons acting in concert to stimulate a burst of TSH secretion from the pituitary gland. This hypothesis is supported by studies that reported disappearance of TSH pulses in

FIGURE 5.13. Serum thyroid-stimulating hormone (TSH) levels measured every 15 minutes in a healthy subject (upper panel), in two subjects with primary hypothyroidism (middle panel) and in a subject with hypothyroidism due to a craniopharyngioma (lower panel). Significant TSH pulses were located by Cluster analysis, a computerized pulse detection program, and are indicated by asterisks (unpublished observations).

hypothyroid rats following administration of TRH antiserum, although TSH levels were near the assay detection limits and therefore TSH pulses may have been obscured [210]. Preliminary data in humans showing that constant TRH infusions do not change TSH pulse frequency cast some doubt on this theory [211]. Endogenous dopamine pathways do not appear to control pulsatile or circadian TSH

Chapter 5

secretion; although dopamine suppresses TSH pulse amplitude, it does not alter TSH pulse frequency. In addition, dopaminergic tone is actually higher at night, and therefore cannot account for the nocturnal TSH surge [212,213]. An alternate explanation for circadian TSH secretion is the diurnal variation in the activity of anterior pituitary 5¢monodeiodinase noted in the rat [214]. However, this has not been confirmed in the human. Recent studies in humans suggest that physiologic serum cortisol levels may control circadian TSH secretion. Subjects with adrenal insufficiency were studied under conditions of short-term glucocorticoid withdrawal and during intravenous administration of hydrocortisone at physiologic levels given as pulses designed to mimic normal pulsatile and circadian cortisol secretion. Daytime TSH levels were higher during glucocorticoid withdrawal, and were decreased by physiologic hydrocortisone infusions, thereby reestablishing the normal TSH circadian rhythm. Twenty four-hour TSH levels were decreased to a similar extent by hydrocortisone infusions at the same dose given as pulses of constant amplitude throughout the 24-hour period, but there was no circadian variation in TSH levels [215]. In parallel studies in healthy subjects, metyrapone (an inhibitor of endogenous cortisol synthesis) decreased endogenous serum cortisol levels during the day, and increased serum daytime TSH levels, leading to abolition of the usual circadian variation in serum TSH levels [216]. These data suggest that the early morning increase in endogenous serum cortisol levels in healthy subjects contributes to the normal daytime decrease in serum TSH levels and leads to the observed normal circadian variation in TSH.

Regulation of TSH Secretion The maintenance of normal TSH secretion depends on complex interactions between central and peripheral hormones (Fig. 5.14). The dominant hypothalamic effect on TSH secretion is stimulatory, via TRH, but there are additional positive and negative hypothalamic modulators of TSH secretion. The dominant peripheral effect on TSH secretion is inhibitory, via thyroid hormone feedback, but other circulating factors modulate TSH secretion. Hypothalamic Control of TSH Secretion

TRH TRH plays a dominant role in the posttranslational processing and release of TSH from the pituitary gland [217,218]. TRH effects on TSH transcription and glycosylation are discussed above (see pp. 179 and 182). Direct TRH effects on TSH secretion occur in vivo and in vitro at concentrations that exist in the pituitary portal blood [134,218,219], and immunoneutralization of TRH in animals leads to a decline in thyroid function [220]. The effects of TRH on TSH processing and secretion are modulated by thyroid state, as discussed in other sections [218] (see p. 187). Immunoreactive TRH is widely distributed throughout the hypothalamus, but the highest concentrations occur in

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185

FIGURE 5.14. Neuroendocrine and peripheral control of thyroid-stimulating hormone (TSH) secretion. T4 , thyroxine; T3 , triiodothyronine; TRH, thyrotropin-releasing hormone; SRIH, somatostatin; DA, dopamine.

the median eminence, preoptic suprachiasmatic nuclei, and paraventricular nuclei (PVN) [221–223]. The latter region is the site of the “thyrotropic area” of the hypothalamus [218]. TRH-containing neurons arise from a group of cells located in the PVN, and project neuronal processes to the median eminence. Lesions of the PVN decrease circulating TRH and TSH levels in normal or hypothyroid animals and cause hypothyroidism [224,225], while electrical stimulation of this area causes TSH release. It is important to note that although baseline levels of TSH are reduced in animals with lesions of the PVN, TSH levels still show appropriate responses to changes in circulating thyroid hormone levels. Thus, the function of TRH in pituitary–thyroid physiology may be to determine the set point of feedback control by thyroid hormones [218]. TRH exerts its effects on TSH secretion via specific high-affinity membrane receptors present on thyrotropes [129,130]. TRH binding to the receptor initiates a cascade of intracellular events that leads to hormone secretion [226,227]. These events have been most clearly elucidated

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in cloned GH3 somatomammotrope cells that secrete prolactin in response to TRH [129], although it appears that TRH-stimulated TSH secretion has a similar mechanism of action [132]. In GH3 cells, the TRH-receptor complex interacts with a guanine nucleotide binding regulatory protein (G) that then binds GTP, causing its activation (G¢). G¢ binds to phospholipase C (C) and activates it (C¢). C¢ catalyzes the hydrolysis of phosphatidylinositol 4,5 bisphosphate (a minor lipid in the plasma membrane), which results in the formation of two intracellular “second messengers,” inositol triphosphate (InsP3) and 1,2-diacylglycerol (1,2DG). InsP3 diffuses from the cell surface membrane to the endoplasmic reticulum, where it causes the release of sequestered Ca2+. This leads to a rapid elevation in intracellular Ca2+ levels, which activates the movement of secretory granules to the cell surface and their exocytosis, either directly or via activation of protein kinase. Simultaneous with these events, there is a parallel activation of protein kinase C by 1,2-DG that also leads to phosphorylation of proteins involved in exocytosis. The prompt and direct effects of TRH on TSH secretion form the basis for the clinical use of exogenous TRH [218]. Acute intravenous administration of 15–500 mg of TRH to normal human subjects causes a dose-related release of TSH from the pituitary. This response is detectable within 5 minutes after TRH administration, and is maximal at 20 to 30 minutes. Serum TSH levels return to basal levels by 2 hours. These normal responses have been compared to TSH levels following TRH administration in patients with primary thyroid disease or hypothalamic–pituitary disease (see p. 196). In contrast to acute bolus injections of TRH, prolonged (2–4-hour) infusions of TRH lead to biphasic increases in serum TSH levels in humans and animals [228]. It is thought that the early phase reflects the release of stored TSH within the thyrotrope, while the later phase reflects release of newly synthesized TSH. Interpretation of TSH responses to more prolonged TRH infusions (24 hours or longer) is complicated by the eventual increase in serum T3 levels, which feedback to suppress further TSH release. This is illustrated by a recent study of 48-hour TRH infusions to intact and hypothyroid-replaced subjects [211]. At all doses of TRH tested, intact subjects had attenuated increases in TSH levels compared to hypothyroid-replaced subjects. However, it should be noted that continuous TRH administration in vitro also causes desensitization of TSH responses, suggesting that increased T3 feedback does not completely explain decreased TSH levels with long-term TRH exposure [229]. SRIH The neuropeptide SRIH is the major hypothalamic inhibitor of GH release. In addition, studies in humans and animals have shown that SRIH inhibits basal and TRHstimulated TSH secretion in vivo and in vitro at concentrations that exist in the pituitary portal blood [230,231]. SRIH also inhibits TRH release from hypothalamic cultures, and SRIH antibodies stimulate TRH release. Therefore,

SRIH may have additional inhibitory effects on TRH secretion at the hypothalamic level. In humans, SRIH infusions suppress TSH pulse amplitude by up to 70%, slightly decrease TSH pulse frequency, and abolish the nocturnal TSH surge [232]. Immunoneutralization of SRIH in animals increases basal TSH levels and TSH responses to TRH [233–235]. Thus, TSH secretion is probably regulated by the hypothalamus through a simultaneous dual control system of TRH stimulation and SRIH inhibition [236]. Immunoreactive SRIH is widely distributed throughout the nervous system and other body tissues, where it exerts a variety of inhibitory actions. In the hypothalamus, the highest concentrations of SRIH occur in the anterior paraventricular region, especially the preoptic, suprachiasmatic and retrochiasmatic nuclei [236]. From these areas, axonal processes of SRIH-containing neurons project to the median eminence. Animals that have undergone sectioning of these fibers have depletion of the SRIH content of the median eminence and increased serum TSH levels [237]. SRIH is secreted in two principal forms: a 14-amino acid peptide, and an N-terminal-extended 28-amino acid peptide. Both of these forms are secreted into the pituitary portal blood in physiologically relevant concentrations [238,239]. It appears that the two forms exert equipotent effects on TSH release [240]. SRIH acts by binding to specific, high-affinity receptors in the anterior pituitary gland [241]. Five subtypes of the SRIH receptor (SSTR1–5) have been identified [242] and SSTR2 and 5 have been localized to thyrotropes [243,244]. Binding of SRIH to its receptor leads to inhibition of adenylate cyclase via the inhibitory subunit of the guanine nucleotide regulatory protein [245], which in turn lowers protein kinase A activity [243] and decreases TSH secretion. SRIH may also exert some effects by cAMP-independent actions on intracellular calcium levels [246]. In addition, SRIH has an indirect suppressive effect on TSH secretion at the pituitary level by decreasing TRH receptors [247]. Other studies had shown that hypothyroidism reduced the efficacy of SRIH in decreasing TSH secretion from bovine pituitaries, and this was reversed by thyroid hormone administration [248]. This was recently explained by studies in mouse thyrotropic tumors which showed that both SSTR2 and 5 were markedly downregulated by the hypothyroid state and were induced by thyroid hormone [244]. Although short-term infusions of SRIH lead to pronounced suppression of TSH secretion, long-term treatment with SRIH or its long-acting analogue octreotide does not cause hypothyroidism [249]. This probably reflects compensatory mechanisms in the thyroid hormone feedback loop. GH deficiency is associated with increased TSH responses to TRH, while GH administration decreases basal and TRH-stimulated TSH secretion [250]. These effects are explained by either GH stimulation of hypothalamic SRIH release or GH mediated increases in peripheral T4 to T3 conversion.

Chapter 5

Dopamine The CNS catecholamine dopamine is a second hypothalamic inhibitor of TSH secretion. Studies in humans and animals have shown that dopamine inhibits basal and TRH-stimulated TSH secretion in vivo and in vitro at concentrations that exist in the pituitary portal blood [251–256]. The administration of dopamine antagonists, including those that do not penetrate the blood–brain barrier, increases TSH levels [257]. In humans, dopamine infusions rapidly suppress TSH pulse amplitude by up to 70%, do not affect TSH pulse frequency, and abolish the nocturnal TSH surge [232]. In addition to acute inhibitory effects on TSH secretion, dopamine also decreases levels of basal and TRH-stimulated TSH subunit gene transcription [131]. Dopamine also has direct effects on hypothalamic hormone secretion that may impact on TSH secretion. For example, dopamine and dopamine-agonist drugs stimulate both TRH and SRIH release from rat hypothalami [258]. However, the net effect of these neurohormone interactions on TSH secretion is unclear. The dopamine that reaches the anterior pituitary gland through the pituitary portal vessels is secreted by neurons whose cell bodies lie in the arcuate nucleus [134]. From the arcuate nucleus, neuronal processes project to the median eminence. Dopamine acts by binding to specific type 2 dopamine receptors (DA2) on thyrotrope cells [259]. This leads to inhibition of adenylate cyclase, which decreases the synthesis and secretion of TSH. In addition, in vitro studies using rat anterior pituitary cells suggest that TSH may down-regulate its own release through the induction of DA2 receptors on thyrotrope cells [260]. The inhibitory effects of dopamine on TSH secretion vary according to sex steroid and thyroid hormone status. For example, dopamine antagonist drugs cause greater increases in serum TSH levels in women than in men [256]. Similarly, dopamine inhibition of TSH release is greater in patients with mild hypothyroidism than in normal subjects, although subjects with severe hypothyroidism may be less responsive [217]. Although short-term infusions of dopamine lead to pronounced suppression of TSH secretion and measurable decreases in serum T3 levels, long-term treatment with dopamine agonists does not cause hypothyroidism. This probably reflects compensatory mechanisms in the thyroid hormone feedback loop. Adrenergic Effects a-adrenergic activation stimulates TSH release directly from the rat pituitary gland at physiologically relevant concentrations of catecholamines [261– 263]. a-adrenergic agonists injected systemically or into the third ventricle stimulate TSH release in rats, while blockade of norepinephrine synthesis or treatment with adrenergic receptor blockers decreases TSH [134,264–266]. It is unclear whether these effects are mediated via changes in TRH and/or SRIH levels, although one study reported that norepinephrine released TRH from hypothalamic cultures. In humans, there are limited data regarding adrenergic effects on TSH secretion. a-adrenergic blockade with phentolamine, which does not readily cross the blood–brain

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barrier, diminishes the serum TSH response to TRH [98]. Administration of thymoxamine, an a-adrenergic blocker that penetrates the blood–brain barrier, causes similar effects [98]. However, administration of epinephrine does not alter TRH-stimulated TSH secretion [267]. These data suggest that endogenous adrenergic pathways have a minor direct stimulatory role in TSH secretion. Noradrenergic stimulation of TSH secretion is mediated by high-affinity a1-adrenoreceptors linked to adenylate cyclase [98]. Therefore, it is likely that dopamine and epinephrine exert opposing actions on thyrotropes by opposite effects on cAMP generation. Opioids In rats, systemic opiate administration suppresses basal or stimulated TSH levels, and the opioid receptor antagonist naloxone reverses these effects [268–271]. In contrast, previous studies in humans have shown that acute opiate administration may slightly stimulate TSH levels, while acute naloxone administration has little effect [272–279]. These findings have supported the statement that opioids have little physiologic role in TSH secretion in humans. However, a recent study has cast doubt on this conclusion [280]. In this study, naloxone was given as a lowdose infusion over 24 hours, and TSH levels were measured frequently. Daytime TSH secretion was preserved, but 24hour TSH secretion decreased by 30%, primarily due to a 40% decrease in nocturnal TSH pulse amplitude. TSH responses to TRH were decreased, suggesting that endogenous opioids stimulate TSH secretion at the pituitary level. Serum T3 levels were decreased as well, suggesting that the magnitude of TSH suppression was sufficient to affect thyroid gland function. If confirmed, these findings suggest that endogenous opioids play an important role in tonic stimulation of TSH secretion. Other Neurotransmitters In addition to the neurotransmitters discussed above, serotonin, g-aminobutyric acid, adenosine, neurotensin, vasoactive intestinal peptide, bombesin, vasopressin, oxytocin, substance P, and cholecystokinin have all been reported to alter TSH secretion [134,281–288]. However, most of these studies have been carried out in animals or in vitro, and the physiologic relevance of these findings to TSH secretion in humans is unknown. Feedback Control of TSH Secretion

The thyroid hormones T4 and T3 exert powerful, dosedependent negative feedback control over TSH synthesis directly at the pituitary level, leading to decreased TSH production (see p. 178). In addition, thyroid hormones acutely block pituitary secretion of TSH [289]. Acute administration of T3 rapidly suppresses TSH levels within hours, while chronic administration leads to further suppression [290,291]. Slight changes in serum thyroid hormone levels within the normal range (induced by thyroid hormone, antithyroid drug or iodine administration) alter basal and TRH-stimulated TSH levels, confirming the sensitivity of

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the pituitary gland to minor alterations in thyroid hormone feedback. Thyroid hormones appear to alter tonic TSH secretion and TSH pulse amplitude without affecting pulse frequency, since subjects with primary hypothyroidism have a near-normal number of TSH pulses of increased amplitude, and T4 replacement in these subjects leads to a decrease in TSH pulse amplitude without much change in pulse frequency [201,207]. Both T4 and T3 function as potent feedback hormones on the thyrotrope. The intracellular monodeiodination of T4 to T3 is greater in the pituitary gland than in peripheral tissues, and contributes approximately half of the intrapituitary content of T3 [292]. Therefore, intrapituitary T4 monodeiodination to T3 may be the mechanism by which the thyrotrope responds to circulating T4 levels. Within the pituitary gland, T3 binds to specific nuclear receptors. The activated T3-receptor complex then binds to specific nucleotide sequences on the TSH subunit genes, inhibiting transcription (see p. 178). In addition to direct effects on TSH gene transcription, thyroid hormones have other actions at the pituitary level that impact on TSH secretion. For example, thyroid hormones decrease the number of TRH receptors [293–295] and stimulate the activity of the pituitary TRH-degrading enzyme [296]. These two effects may act in concert to decrease TRH stimulation of TSH secretion. Thyroid hormones exert direct effects on hypothalamic function that modulate TSH secretion. TRH mRNA levels in the PVN are increased in hypothyroidism and are reduced by T3 and T4 [297–300]. Hypothalamic lesions in the PVN prevent the full increase in plasma TSH and pituitary TSH b mRNA levels seen in response to hypothyroidism [301]. Hypothalamic SRIH content is decreased in hypothyroid rats, and is restored by T3 treatment. Finally,T3 directly stimulates SRIH release from hypothalamic tissue [302]. These combined effects of thyroid hormones on TRH and SRIH decrease TRH release from the hypothalamus, and thereby indirectly decrease TSH secretion. Other Peripheral Factors that Affect TSH Secretion

Glucocorticoids Pharmacologic doses of glucocorticoids suppress basal TSH levels, blunt TSH responses to TRH, and attenuate or abolish the nocturnal TSH surge in humans and animals [303–311]. Some patients with Cushing’s syndrome have low TSH levels and loss of the nocturnal TSH surge, probably due to endogenous hypercortisolism. In healthy subjects, infusions of hydrocortisone (100–300 mg/day) that increase serum cortisol to levels seen in moderate stress suppress 24-hour TSH secretion by 60% to 70% [312]. Infusions of much lower doses of hydrocortisone in healthy subjects (34–46 mg/day) lead to similar decrements in 24hour TSH secretion. These hydrocortisone doses were designed to lead to mild increases in serum cortisol levels, similar to endogenous levels seen during short-term fasting or other minor stressors. The resulting serum cortisol levels, although higher than baseline endogenous levels, are still

within the “normal” range for healthy, nonstressed subjects. These data suggest that even low-dose, near-physiologic glucocorticoid administration affects TSH secretion, and that mild increases in endogenous cortisol levels may exert significant effects on TSH levels [313]. In all of these studies, the changes in TSH levels are due to decreased TSH pulse amplitude without alteration in TSH pulse frequency, with more profound suppression of nocturnal TSH secretion and abolition of the TSH surge. Physiologic glucocorticoid levels also affect TSH secretion [215,216,304,306,314,315]. Many reports have described elevated serum TSH levels in untreated patients with adrenocortical insufficiency that resolve with steroid replacement. These studies are compromised by the high incidence of autoimmune thyroid disease in these patients, since glucocorticoids could have independent immunomodulating effects on autoimmune thyroid disease. However, studies of metyrapone (an inhibitor of cortisol synthesis) administration to healthy subjects have confirmed a reciprocal relationship between endogenous cortisol and TSH levels [216], and studies of physiologic hydrocortisone replacement in patients with adrenal insufficiency show that increased daytime TSH levels seen in the absence of cortisol are decreased by physiologic cortisol levels [215]. Glucocorticoid suppression of TSH levels may occur directly at the pituitary gland, and/or via hypothalamic effects. The former possibility is supported by the maintenance of TSH pulse frequency and the suppression of TSH responses to exogenous TRH after glucocorticoid administration [312]. Some animal studies suggest that glucocorticoids exert direct effects on thyrotropes, although they appear to be highly dependent on dose and time-course of steroid administration [316–319]. However, in contrast to effects of thyroid hormones, it does not appear that glucocorticoids directly affect TSH gene transcription. For example, administration of dexamethasone to rats does not alter pituitary TSH content or TSH b-subunit mRNA levels, although it does attenuate TSH elevations and enhance T3-mediated TSH suppression in hypothyroid rats [139,320]. In addition to direct pituitary effects, other studies have shown that dexamethasone increases hypothalamic TRH levels, while circulating TRH levels are decreased. These results suggest that dexamethasone may inhibit TRH release from the hypothalamus [321]. Reports that the proposed consensus sequence for glucocorticoid receptor binding is present in the 5¢-flanking region of the TRH gene also support the possibility of hypothalamic effects of glucocorticoids on TRH [322]. Patients with Cushing’s syndrome or subjects receiving prolonged courses of glucocorticoids may have low serum T4 as well as TSH levels. Whether such patients have tissue hypothyroidism, and whether they should be treated with thyroid hormone, is unclear; however, patients with acute or chronic illnesses and similar abnormalities in thyroid hormone levels do not appear to benefit from thyroid hormone therapy.

Chapter 5

Sex Steroids Serum and pituitary TSH concentrations are higher in male than in female rats, are reduced by castration, and can be restored by androgen administration [54]. Exogenous testosterone administration to castrated male or female rats increases basal and TRH-stimulated serum TSH levels [323]. In contrast, androgen administration to intact female rats decreases pituitary TSH b-subunit mRNA concentrations without altering serum or pituitary levels of TSH [140]. Estrogen administration to euthyroid rats does not alter serum TSH levels. In contrast, in hypothyroid rats, pharmacologic doses of estrogen may enhance thyroid hormone feedback of TSH synthesis. In euthyroid humans, early studies reported that androgens suppress TSH responses to TRH. However, most studies suggest that changes in endogenous or exogenous sex steroid levels do not significantly affect basal or TRHstimulated TSH levels [324,325]. Other studies confirm that there is no significant gender difference in the basal mean and pulsatile secretion of TSH or in the TSH response to TRH [326]. Therefore, although the data are not conclusive, sex steroids do not appear to play a major regulatory role in TSH secretion under normal conditions in humans. Cytokines The cytokines, which are circulating mediators of the inflammatory response that are produced by many cells, have been shown to affect the hypothalamic– pituitary–thyroid axis. Tumor necrosis factor (TNF) and interleukin-6 (IL-6) decrease serum TSH levels in healthy human subjects, and TNF and interleukin-1 (IL-1) decrease TSH levels in animals [168,327–331]. Administration of these cytokines leads to thyroid hormone levels similar to those seen in acute nonthyroidal illness [332]. In rats, TNF reduces hypothalamic TRH content and pituitary TSH gene transcription. IL-1 stimulated type II 5¢-deiodinase activity in rat brain, which may decrease TSH secretion by increasing intrapituitary T3 levels. In addition, IL-1 and IL-6 are produced by rat anterior pituitary cells, and may serve as autocrine or paracrine factors in TSH secretion during illness [333,334]. Nonsteroidal Anti-inflammatory Agents The nonsteroidal anti-inflammatory agents (NSAIDs), including aspirin, salsalate, ibuprofen, indomethacin, and naproxen, are used extensively to treat a wide variety of acute and chronic inflammatory conditions. Their major mechanism of action appears to be inhibition of prostaglandin synthesis. A number of studies suggest that many NSAIDs lead to decreased serum TSH levels, within hours of administration [335–340]. Usually TSH levels are lower than nontreated baseline, but remain within the broad normal range. However, serum TSH levels can occasionally decrease below the normal range in patients taking NSAIDs, especially at higher doses. In most cases, this decrease in TSH is thought to be an indirect effect due to NSAID displacement of thyroid hormones from protein binding sites, with transient increase in free thyroid hormone levels and feedback sup-

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pression of TSH. However, one study suggests that at least one of the NSAID classes (meclofenamate and fenclofenac) may directly inhibit TRH-induced TSH release by rat anterior pituitary cells in vitro [341]. ACTION OF TSH TSH acts on the thyroid gland by binding to the TSH receptor [342]. Excellent reviews of this subject have been published [100,343,344]. This receptor is located on the plasma membranes of thyroid cells and consists of a long extracellular domain, a transmembrane domain and a short intracellular domain. Knowledge of the molecular structure of the receptor has allowed a better understanding of the mechanism of action of TSH that results in the production of thyroid hormone. Studies with recombinant TSH receptor have provided insights into the role of TSH receptor antibodies in autoimmune thyroid disease. In addition, mutations of the TSH receptor have been described that result in clinical syndromes of hypothyroidism and hyperthyroidism.

TSH Receptor Gene The human TSH receptor gene is located on chromosome 14 locus q31 and spans a region greater than 60 kb in size containing ten exons [345]. Exons one through nine have 327, 72, 75, 75, 75, 78, 69, 78, and 189 bp, respectively, and encode part of the extracellular domain, whereas exon ten is greater than 1412 bp and encodes the rest of the extracellular domain as well as all of the transmembrane and the intracellular domains. The promoter region of the human TSH receptor gene has also been partially characterized [345]. The major transcriptional start site, designated as +1, is located 157 bp upstream of the translation initiation codon ATG. There are no consensus CAAT or TATA boxes but there are degenerate CAAGGAAAGT and TAGGGAA boxes located at positions -86 and -43, respectively. The regions of the promoter important for tissue-specific expression and those responsive to TSH and cAMP, the two main regulators that have been shown to inhibit the rat TSH receptor gene expression [346], are yet to be defined. Northern blot analysis has revealed two major transcripts of the human TSH receptor of 3.9 and 4.6 kb in size that differ only in the length of the 3¢ untranslated region [347,348].

TSH Receptor Structure The TSH receptor is synthesized as a single polypeptide chain of 764 amino acids that includes a 20-amino acid signal peptide [349]. However, the TSH receptor has been found to exist on the cell surface as a single chain and also as a two-subunit form, produced by internal cleavage apparently at two sites, releasing a potentially immunogenic 5– 7-kDa peptide [344,350]. The amino-terminal half of the protein contains 16 hydrophilic leucine-rich repeats (LRR)

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that form the extracellular domain and includes six potential glycosylation sites. The asparagine-linked oligosaccharides appear to be important for correct folding, membrane targeting and receptor function [351,352]. The LRR are the common feature of the superfamily of LRR proteins. One of these, the ribonuclease inhibitor, has been cocrystallized with its ligand [353], and this has allowed the construction of a model of the extracellular domain of the TSH receptor bound to TSH [354], as shown in Fig. 5.15. The carboxyl-terminal half of the protein, modeled after the structure of the G protein-coupled receptors, based on rhodopsin [355], is also shown in Fig. 5.15. This region contains seven hydrophobic transmembrane segments, three extracellular loops, three cytoplasmatic loops and a short cytoplasmatic tail of 82 amino acids (Fig. 5.15).

Determinants of TSH Interaction with its Receptor The entire extracellular domain as well as parts of the transmembrane domain of the TSH receptor contribute to TSH binding. However, two regions, from residue 201 to 211 and 222 to 230, are particularly important in TSH-specific binding [356]. In contrast, a different region of the extracellular domain, from residue 287 to 404, appears to be important for binding of TSH receptor antibodies in autoimmune thyroid disease [357]. Other authors have reported considerable overlap of TSH binding regions and antibody epitopes [343]. Interestingly, studies using rat FRTL-5 cells have shown that thyroid stimulating autoantibodies (TSAbs or TSI) stimulate whereas TSH binding inhibitory antibodies (TBIAbs) inhibit TSH-mediated gene expression, suggesting that these antibodies must act on different epitopes of the receptor that differ in their signal transduction mechanism [346]. The transmembrane domain of the TSH receptor also appears to be important in ligand binding. A point mutation in the fourth transmembrane domain of the TSH receptor gene has been described in the hyt/hyt congenitally hypothyroid mouse that abolishes TSH binding [358]. Inactivating TSH receptor mutations are a rare cause of congenital hypothyroidism in humans, although 12 such mutations have been identified, eight of which map to the aminoterminal domain [359–362]. Specificity of TSH binding is conferred by the TSH b-subunit. It appears that amino acid residues from 58 to 69, within the bL3 loop, and from 88 to 105, the “seat-belt” region of the TSH b-subunit [363] play an important role in binding to and activation of the TSH receptor. The carboxylterminal end of TSH b contains multiple lysine residues (positions 101, 107 and 110) and a cysteine at position 105 that are critical for the ability to bind to the receptor [364]. Congenital hypothyroidism due to biologically inactive TSH was found to result from a frameshift mutation with loss of b-cysteine105 [365,366] (see p. 197). Several regions of the a-subunit are also important for TSH activity, particularly the residues a11–20 and a88–92 (see Fig. 5.12)

FIGURE 5.15. Schematic model of the human thyroidstimulating hormone (TSH)–TSH receptor complex. The receptor is depicted in accordance with models based on the leucine rich repeats (LRR)-containing ribonuclease inhibitor (353) and G proteincoupled rhodopsin (355). In the center, the a-subunit (white ribbon) and TSHb-subunit (black ribbon) are shown folded and combined as depicted in Fig. 5.12, except that the TSH molecule is inverted, so that the a hairpin loops are oriented toward the extracellular loops of the trasmembrane domain of the receptor, and the b hairpin loops toward the concave surface of the LRR. From Grossmann et al. [100].

[176,367]. In addition, the oligosaccharide chain at position a-asparagine52 plays an important role in both binding affinity and receptor activation. A mutant TSH lacking the a-asparagine52 oligosaccharide showed increased in vitro activity, although this same mutation had the opposite effect on CG binding to its native receptor [368]. However, such a mutation also increased TSH clearance and this decreased in vivo activity [368]. In addition, the oligosaccharide chains on the TSH subunits are critically important for signal transduction [149,150,369]. In this regard, the a-subunit oligosaccharides are important for all the pathways activated by the receptor, whereas the TSH b-subunit oligosaccharide only

Chapter 5

influences the adenylate cyclase pathway [370]. The mechanism by which the oligosaccharides influence signal transduction is not known. A model for the action of the glycoprotein hormones has been proposed that suggests a role for the oligosaccharides in directly modulating the influx of calcium into the target cell [371]. The ability of chorionic gonadotropin to bind to the TSH receptor was demonstrated in rat thyroid cells [372] and recently confirmed in studies using recombinant human TSH receptor [373,374]. The activity of CG was estimated to be less than 0.1% compared to TSH. LH was found to have a 10-fold higher potency for activation of the TSH receptor when compared to CG, but a mutant of CG that lacks the carboxyl-terminal region of CG b from amino acid residues 115 to 145 showed a potency equivalent to that of LH [374]. This truncated form of CG is one of the forms in the heterogeneous population of CG molecules produced in normal pregnancy and in trophoblastic tumors, and may be present in amounts sufficient to cause significant thyroid gland stimulation [375]. The ocurrence of gestational hyperthyroidism due to a mutation in the TSH receptor that increases its sensitivity to CG has also been described [376].

Signal Transduction at the TSH Receptor The three intracytoplasmatic loops of the transmembrane domain appear to be important for signal transduction [377]. The TSH receptor is coupled to the Gs protein cascade, probably through the carboxyl-terminus of the third cytoplasmatic loop [378]. Thus, binding of TSH activates adenylate cyclase to produce cAMP [379,380]. The phosphatidylinositol pathway is also activated [381], but this pathway is slower and requires a higher concentration of TSH [382,383]. The unliganded TSH receptor has been found to have significant constitutive activity [384,385], suggesting that regulation may involve the release of an inhibitory restraint. This would explain the relatively high frequency of activating mutations of the TSH receptor compared to inactivating mutations. In cases of congenital hyperthyroidism [384,386], the mutations were located in the extracellular domain and the second, fourth, fifth and sixth transmembrane domains, while in the hyperfunctioning adenomas [384,387] the mutations were found to localize to the carboxyl-terminus of the third cytoplasmatic loop and adjacent sixth transmembrane domain. All these mutations resulted in constitutive activation of adenylate cyclase [388]. Germline mutations of the cytoplasmatic tail of the TSH receptor have been described in 33.3% of patients with toxic multinodular goiter and 16.3% with Graves’ disease, and this mutation was found to result in an exaggerated cAMP response to TSH [389]. In addition, specific amino acids in the third cytoplasmatic loop have been identified that are important for the phosphatidylinositol pathway but do not appear to play a role in the adenylate cyclase pathway [390].

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Effects of TSH TSH action on the receptor results in activation of the adenylate cyclase pathway and to some extent the phosphatidylinositol pathway, as described above, and leads to the activation of protein kinase A and C with resulting phosphorylation of multiple proteins, including nuclear histones [391]. Proteins phosphorylated by the protein kinase C pathway appear to be different from those phosphorylated by protein kinase A [392]. In addition, phosphoprotein phosphatases are activated and lead to the dephosphorylation of another set of proteins [392]. The effects of TSH on the thyroid gland include changes in cell morphology, iodine metabolism, synthesis of thyroid hormone and thyroid gland growth. Effects of TSH on Thyroid Cell Morphology

TSH causes dramatic changes in the morphology of the thyroid. The initial response to TSH is the incorporation of exocytotic vesicles into the cell membrane at the apical pole of the follicular cells that is quickly followed by formation of cytoplasmatic projections and microvilli [393]. The number of cytoplasmatic projections has been correlated with the level of TSH [394]. After 8 hours of stimulation, the follicular cells become columnar and filled with colloid droplets and luminal colloid is nearly depleted collapsing the follicles [395]. In contrast, when TSH decreases the follicular cells become cuboidal and colloid increases, distending the follicles [395]. Lysosomes migrate from the basal pole toward the apical pole where they fuse with the colloid droplets and then migrate toward the basal pole, becoming smaller and denser [393]. The cytoskeletal system, that includes myosin, actin, tropomyosin, calmodulin, profilin and tubulin, has been implicated in this process [396,397]. Effects of TSH on Iodine Metabolism

Iodide transport into the cell is stimulated by TSH via the adenylate cyclase pathway [398]. Although thyroid peroxidase transcription and mRNA stability are increased by TSH through the adenylate cyclase pathway [399], the generation of peroxide and iodide organification appear to be mediated by a phosphatidylinositol pathway independent of protein kinase C [400]. Conversely, iodine inhibits activation of the adenylate cyclase system by TSH [401]. Effects of TSH on the Synthesis of Thyroid Hormone

The endpoint of TSH action is the production of thyroid hormone by the thyroid gland. The process begins with thyroglobulin gene transcription. The transcriptional rate and possibly the mRNA stability are increased by TSH [402]. TSH stimulates iodide uptake and organification, as described above. TSH then acts on the iodinated thyroglobulin stored in the luminal colloid and stimulates its hydrolysis resulting in the release of the constituent amino acids, including the iodotyronines T3 and T4.

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Effects of TSH on Thyroid Gland Growth

After long-term stimulation by TSH, the thyroid gland enlarges as a result of hyperplasia and hyperthophy. Acutely, TSH has a rapid mitogenic effect on the thyroid gland that is evident within 5 minutes [403]. It increases DNA synthesis through the adenylate cyclase pathway [404], specifically through activation of protein kinase A type I [405]. TSH may also regulate growth by cAMP-independent pathways [406] and interactions with the action of the growth factors epidermal growth factor (EGF) and insulinlike growth factor-I (IGF-I) [407,408]. It has been found that TSH and other growth factors increase the mRNA levels of the proto-oncogenes c-fos and c-myc in rat thyroid cells [409]. TSH inhibits apoptosis [410], perhaps by regulating p53 and bcl-2, as shown for gonadotropins [411,412]. Mutations within the a33–44 region were found to reduce growth stimulation but not affect cAMP production [413], although a clear dissociation of the various actions by TSH analogs has not yet been achieved. TSH-induced Receptor Desensitization

The phenomenon of desensitization has been described both in vivo and in vitro, whereby prior TSH stimulation leads to a 30% to 70% decrease in the subsequent cAMP response to TSH stimulation [414,415]. Recent studies using recombinant TSH receptor have shown that desensitization does not occur when the receptor is expressed in nonthyroidal cells, suggesting that this phenomenon requires a cellspecific factor [416]. Extrathyroidal Actions of TSH

The ocurrence of precocious puberty in cases of severe juvenile primary hypothyroidism [417] has suggested that high levels of TSH are able to cross-activate the gonadotropin receptors. This interaction has now been demonstrated using recombinant human TSH, which has been found to be capable of activating the FSH [418] but not the CG/LH receptor [419]. Expression of both TSH and its receptor has been reported in lymphocytes [420,421], and has led to the speculation that TSH may have other nonclassic functions. The TSH receptor was also found to be expressed in adipocytes [422], and TSH was found to stimulate proliferation and inhibit differentiation of preadipocytes in vitro [423]. More studies are needed to determine the physiological significance of the extrathyroidal effects of TSH, as this may impact the safety of future treatment modalities for thyroid cancer that may attempt to target radioisotopes to the TSH receptor [424,425].

TSH MEASUREMENTS Accurate and specific measurements of serum TSH concentrations have become the cornerstone for diagnosing and treating the vast majority of thyroid disorders. The first mea-

surements of TSH in circulating blood used insensitive and nonspecific bioassays that measured an end point of TSH action, for example, a rise in cyclic AMP levels in cultured thyroid cells [426–428]. Such assays were labor-intensive and time-consuming. Subsequently, immunological approaches were used to measure TSH in human blood when antibodies to human TSH were first developed in 1963. The availability of these antibodies and crudely purified human TSH preparations allowed investigators to develop the first radioimmunoassays for human TSH.

Radioimmunoassay of TSH During the two decades between 1963 and 1983 radioimmunoassays were widely employed for the measurement of TSH in human serum [429–434]. Modifications and improvements in the techniques of radioimmunoassays allowed serum TSH assays to be moderately sensitive and specific [435–439]. The most sensitive radioimmunoassays of TSH had functional detection limits between 0.5 to 1 mU/l of TSH. These assays, known as “first generation assays,” were useful in distinguishing elevated serum levels of TSH in primary hypothyroidism from those normal values found in euthyroid subjects. One hundred percent of primary hypothyroid subjects had elevated TSH levels. In contrast, these “first generation assays” could not accurately quantitate values within the normal range, and there was considerable overlap with the values found in euthyroid and hyperthyroid subjects.

Immunometric Assay of TSH The first monoclonal antibody to human TSH was developed in 1982 and introduced a new era in the technology for measuring TSH [440]. Monoclonal antibodies had defined epitope specificity allowing them to be used in “sandwich-type” assays subsequently termed immunometric assays (Fig. 5.16) [441]. Immunometric assays were quickly shown to be superior to classic radioimmunoassays [442– 448]. The principles of these assays are quite different. Radioimmunoassays used low amounts of labeled antigen in combination with very dilute antibody mass. In contrast, immunometric assays label one of the monoclonal antibodies, the “signal antibody.” This signal may be radioisotopic or chemiluminescent. Another monoclonal antibody with a different epitope specificity is linked to a solid support and is called the “capture antibody.” Both antibodies are used in excess, sandwiching TSH between the two antibodies and thus capturing all TSH molecules in a given sample. These modifications in the measurement of TSH resulted in important changes. First, the assays were highly specific with no cross-reaction to the other human glycoprotein hormones. Second, 100% of euthyroid controls have detectable and quantifiable levels of TSH residing within a normal range of approximately 0.5–5 mU/l. Third, there is little or no overlap in TSH values in patients with hyperthyroidism

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FIGURE 5.16. Schematic representation of an immunometric assay for thyroid-stimulating hormone (TSH). The assay uses one labeled monoclonal antibody (the signal antibody) and another monoclonal antibody with a different epitope specificity linked to a phase face support (the capture antibody). All TSH molecules are sandwiched between the two monoclonal antibodies. From Ridgway [441].

FIGURE 5.17. Improved thyroidstimulating hormone (TSH) assays with greater discrimination between normal controls and hyperthyroid subjects. A schematic representation of TSH levels in euthyroid (white circles) vs thyrotoxic (black circles) samples. The sera were measured in first, second, third, and fourth generation TSH assays. The lower limit of sensitivity for each generation of assay is shown by the vertical bars. From Spencer [452].

compared to euthyroid controls. Thus, immunometric TSH assays allow physicians to immediately discriminate euthyroid subjects from both hyperthyroid and hypothyroid patients [449–451]. The degree to which a given assay can separate undetectable TSH levels found in hyperthyroid subjects from normal values in euthyroid controls had improved steadily (Fig. 5.17) [452]. These improvements have resulted in progressively lower functional detection limits, defined as the lowest TSH value detected with an interassay coefficient of variation £20% (Fig. 5.18) [452]. Thus, first generation assays (usually radioimmunoassays) have functional detection limits of 1–2 mU/l, second generation assays 0.1–0.2 mU/l, third generation assays 0.01–0.02 mU/l and fourth generation assays 0.001–0.002 mU/l. At the present time, the most sensitive commercially available TSH assays are third generation assays. The most widely used assays, however, are second generation. Only research laboratories which have extensively modified assay procedures are able to reach fourth generation technical performance [451].

Free TSH b- and a-subunit Measurements In 1974 the TSH b- and a-subunits were purified from human TSH and specific antibodies to them developed [453]. The availability of specific antibodies to free TSH band a-subunit allowed for the development of radioimmunoassays to these subunits. In general, measurements of free TSH b have not been clinically useful because they have been very concordant with measurements of intact TSH [454,455]. In addition, the free TSH b assay is not very sensitive and free TSH b-subunit levels are not detectable in either euthyroid subjects or patients with hyperthyroidism. In contrast, measurement of the free a-subunit has been useful in the evaluation of pituitary and placental disease because in these diseases, there can be discordances between free a-subunit levels and intact hormone levels. Free asubunit is detectable and measurable by radioimmunoassay in euthyroid and eugonadal human subjects [455–458]. More recently, immunometric assays for free a-subunit have

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FIGURE 5.18. Schematic representation of the generation system for thyroid-stimulating hormone (TSH) assay nomenclature. Each generation represents a 10-fold improvement in functional assay sensitivity. The black bars denote the 95% confidence limit of measurement. For example, at TSH levels of 0.01 to 0.02 mU/l with a 20% interassay coefficient of variation, third generation assays have a 95% confidence limit corresponding to the horizontal axis values indicated by the black bar. From Spencer [452].

FIGURE 5.19. Schematic representation of the use of sensitive thyroid-stimulating hormone (TSH) assays in patients with increasing free T4 levels. There is a log/linear relationship between serum TSH concentrations and free T4 concentrations.

been developed which reveal a normal range for free asubunit of 0.1–1.6 ng/mL [459]. Elevated values of free asubunit are found in the sera of patients with TSH-secreting or gonadotropin-secreting pituitary tumors [460–467], as well as choriocarcinoma [468]. Elevated free a-subunit levels have also been reported in a variety of non-pituitary and non-placental malignancies including cancers of the lung, pancreas, stomach, prostate, and ovary [469–476].

Provocative Testing of TSH Basal measurements of TSH and free a- or TSH b-subunits are best interpreted in the context of knowledge of free T4 and/or free T3 serum concentrations. The sensitive inverse and reciprocal negative feedback regulation between circulating free T4 levels and TSH secretion permits accurate diagnoses to be made (Fig. 5.19). However, under certain

circumstances provocative stimulation of TSH is necessary for a full understanding of the integrity of the hypothalamic–pituitary–thyroid axis. This subject has recently been reviewed [477]. The TRH stimulation test was developed to investigate the integrity of this axis. TRH is a hypothalamic tripeptide (pyroglutamyl-histidyl-prolineamide) that directly stimulates TSH biosynthesis and secretion. Given intravenously, intramuscularly or orally, TRH causes a reproducible rise in serum TSH levels in euthyroid subjects [134,218]. The test is performed by giving 200–500 mg of TRH intravenously and measuring TSH at 0, 20–30, 60, 120 and 180 minutes after injection. In euthyroid subjects, there is an immediate release of TSH rising to peak levels approximately 20–30 minutes after TRH injection, usually reaching values five- to 10-fold higher than basal [478,479]. Using third and fourth generation assays across a wide spectrum of normal and abnormal basal TSH levels,

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FIGURE 5.20. The relationship between basal and absolute (thyrotropinreleasing hormone [TRH]-stimulated minus basal thyroid-stimulating hormone [TSH]) TRH-stimulated TSH response in 1,061 ambulatory patients with an intact hypothalamic–pituitary (H–P) axis compared with that in untreated and T4-treated patients with central hypothyroidism. From Spencer et al. [451].

the increase of TSH after TRH stimulation has been between 2.8- and 22.9-fold [451] (Fig. 5.20). In hyperthyroid subjects with undetectable basal serum TSH levels, there is little or no response to TRH stimulation [449, 451]. Patients with low basal serum TSH levels secondary to pituitary insufficiency (secondary hypothyroidism) or hypothalamic disease (tertiary hypothyroidism) have absent or attenuated TSH responses to TRH [480,481]. In contrast, patients with elevated TSH levels due to primary hypothyroidism have exuberant responses to TRH stimulation. However, elevated TSH levels in patients with pituitary TSH-secreting tumors usually fail to respond to TRH stimulation [462] (Fig. 5.21).

DISORDERS OF TSH PRODUCTION Abnormalities in TSH secretion and the integrated circadian rhythms of pulsatile TSH secretion can be caused by diseases in the hypothalamus, pituitary and thyroid gland. In some instances, novel molecular mechanisms have precisely defined the causes of defective TSH secretion. In other instances, descriptive pathophysiology awaits more definitive mechanistic explanations for the disorders.

Hypothalamic Disorders Since the hypothalamus is the source of factors that have important effects on TSH production and secretion, diseases in this part of the brain can dramatically influence serum TSH levels. Theoretically, diseases which affect the production of the hypothalamic factors including TRH, SRIH, or dopamine could influence TSH production. Structural abnormalities in the hypothalamus have resulted in clinical hypothyroidism, which presumably is due to decreased or defective TRH production (Table 5.1). Unfortunately, this pathophysiological explanation for the hypothyroidism is theoretical, since accurate measurements of hypothalamically-derived TRH are not readily available. Measurements of TRH in peripheral plasma are difficult to interpret since the source of TRH in plasma is largely due to extrahypothalamic TRH production [482,483]. Nevertheless, indirect evidence has suggested that TRH deficiency plays an important role in the etiology of hypothyroidism in these cases. Indeed, exogenous administration of TRH to patients with hypothalamic hypothyroidism can restore serum thyroid hormone levels towards normal and result in clinical improvement [484,485]. Elevated SRIH or dopamine could also cause decreased TSH

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FIGURE 5.21. Schematic representation of thyrotropin-releasing hormone (TRH) stimulation tests in patients with a variety of thyroid disorders.

Table 5.1. Neoplasia

Infiltrative

Trauma

Infection

Vascular Congenital

Causes of hypothalamic hypothyroidism Pituitary adenoma Craniopharyngioma Dysgerminoma Meningioma Sarcoidosis Histiocytosis X Eosinophilic granuloma Radiation Head injury Post surgical Tuberculosis Fungus Virus Stalk interruption Midline defects Rathke’s pouch cysts

thus effecting hypothalamic hypothyroidism. Indeed, administration of GH has been associated with this syndrome [250,486], presumably by stimulating hypothalamic SRIH production. However, until measurements of hypothalamic SRIH secretion are available, this mechanism also remains theoretical. In fact, enhanced peripheral conversion of T4 to T3 induced by GH could have the same effect. Future studies involving the use of specific antagonists to the SRIH or dopamine receptors on pituitary thyrotropes may prove useful in unraveling specific causes of hypothalamic hypothyroidism. The serum levels of TSH in hypothalamic hypothyroidism may be either low, normal, or even minimally elevated [480,487–490]. The paradoxical finding of low serum thyroid hormone levels in association with normal or elevated basal TSH levels suggest that the circulating TSH in hypothalamic hypothyroidism is biologically defective [484,485,488]. In fact, TRH deficiency has been associated

FIGURE 5.22. Twenty-four hour thyroid-stimulating hormone (TSH) pulsations measured every 15 minutes in a patient with central hypothyroidism due to hypothalamic hypothyroidism (solid line) vs a normal control (dashed line). The nocturnal surge of TSH is not present in the patient with central hypothyroidism, although the pulse frequency is similar in both the patient and the normal control. From Samuels et al. [208].

with differences in glycosylation patterns of the TSH molecule which result in decreased TSH receptor binding and activation [163,485] (see pp. 182, 185 and 190). The 24hour secretory profile of TSH in patients with hypothalamic hypothyroidism is also abnormal [208,488,491,492]. The frequency of the TSH pulses is the same as euthyroid controls, but the amplitude of the pulses is decreased, particularly at night-time [208], resulting in a loss of the normal nocturnal surge (Fig. 5.22) (see p. 186).

Pituitary TSH Disorders Abnormal pituitary function can be the sole cause for altered TSH secretion. Increases or decreases in pituitary

Thyroid-stimulating Hormone

Chapter 5

opment of thyrotropes, somatotropes, and lactotropes, was found to have mutations in the coding region that alter the function of the Pit-1 protein or completely disrupt its structure. The absence of Pit-1 prevents normal pituitary development resulting in hypoplasia of the pituitary. In heterozygotes, where a normal allele is present, the abnormal Pit-1 protein can bind to DNA but not be able to effect transactivation, interfering with the function of the normal Pit-1 (dominant negative mechanism). This disorder often results in pituitary hypoplasia in addition to the hormone deficiencies. Interestingly, a similar combined hormone deficiency syndrome has been reported in two murine models in which the Pit-1 gene is defective: a point mutation found in the Snell dwarf (dw) and [12] and a major deletion in the Jackson dwarf (dwJ) [499]. A more frequent cause of CPHD has recently been delineated by the discovery of a new pituitary-specific transcription factor called “prophet of Pit-1” (PROP-1). This factor is a paired-like homeodomain protein in which a mutation in the murine species causes the Ames dwarf (df) mouse phenotype [22]. Subsequently, patients with CPHD were found to have mutations in the PROP-1 gene [500–502]. Most recently, 50% of families with CPHD have been shown to contain mutations in the PROP-1 gene [21], far exceeding the prevalence of mutations in the Pit-1 gene as a cause for CPHD. The mutations are all found in the homeodomain part of the molecule. Interestingly, the phenotype of patients with PROP-1 mutations includes deficiencies not only of GH, prolactin and TSH but also of LH and FSH. Furthermore, the development of hormone deficiencies may not be neonatal but rather progressively occur up to the age of adolescence (Fig. 5.23). To date no patient with these mutations has developed clinically relevant ACTH deficiency. Thus, congenital hypothyroidism due to pituitary TSH deficiency has been found at the molecular level to be due to germ cell point mutations of either the pituitary-specific

TSH production can be caused by either congenital or acquired abnormalities. Decreased Pituitary TSH Production–Congenital

Congenital hypothyroidism can be caused by a defective or absent production of TSH. It can present as isolated TSH deficiency or the TSH deficiency may be combined with other anterior pituitary defects. The patients are born with typical clinical features of congenital hypothyroidism, but are remarkable for the absence of a goiter. Circulating levels of free T4 and T3 are low and TSH levels are usually undetectable. In addition, serum TSH levels fail to increase with a standard TRH stimulation test. The disease is generally inherited as an autosomal recessive disorder, and affected individuals have severe mental and growth retardation. Molecular mechanisms which are responsible for the isolated TSH deficiency have been described in a limited number of cases (Table 5.2). In one family a single base substitution at nucleotide position 145 of the TSHb gene altered the CAGYC region [180,493], a critically important contact point for the noncovalent combination of the TSH b-and a-subunits (see p. 183). In another kindred, a single base substitution introduced a premature stop codon, resulting in a truncated TSH b-subunit which included only the first 11 amino acids [148]. In other cases, the disorder involves the production of biologically inactive TSH with loss of bcysteine105 [365,366,494], resulting in a similar phenotype, except that in some of these cases circulating TSH was detectable. Another cause of congenital pituitary TSH deficiency is combined pituitary hormone deficiency (CPHD). Subjects with this disorder have congenital hypothyroidism and growth retardation secondary to TSH, GH and prolactin deficiencies [495–498]. Within the last decade, molecular mechanisms for this unusual defect have been elucidated [16–19]. The pituitary-specific transcription factor Pit-1 (see p. 173), which is critically important for normal devel-

Table 5.2.

197

Congenital hypothyroidism: isolated TSH b defects I

II

III

Inheritance Syndrome Serum T4 Serum TSH Response to TRH Nucleotide change Protein defect

Autosomal recessive Cretinism Ø None detected None detected Missense, G145A G29R (CAGYC region) No combination with a

Autosomal recessive Cretinism Ø Ø Impaired Deletion, T410 C105V and frameshift. (seat-belt region) Inactive TSH

Reported cases

3 families in Japan [180]

Autosomal recessive Cretinism Ø None detected None detected Nonsense, G94T Premature stop (bL1 loop region) Truncated TSHb (11 amino acids) 2 families in Greece [148]

2 families in Brazil [365], 2 families in Germany [366,494]

100

35

GH TSH LH/FSH

80

Percentage

Hypothalamic–Pituitary Function

SECTION 1

28

60

21

40

14

20

7

0

0

2

4

6

8

10

12

14

16

18

Years

transcription factors PROP-1 or Pit-1 or the TSH b-subunit gene. Decreased Pituitary TSH Production–Acquired

The acquired causes of pituitary TSH deficiency generally relate to destructive processes in the anterior pituitary. These can include infiltrative or infectious disorders, compression secondary to neoplastic processes, or active ischemic and hemorrhagic processes involving the pituitary gland. The most common cause of acquired pituitary TSH deficiency is neoplastic destruction of normal anterior pituitary cells by either a primary pituitary neoplasm, a craniopharyngioma, or infiltrating metastatic disease to the pituitary. In acquired pituitary TSH deficiency multiple pituitary deficiencies are concomitantly associated; acquired isolated TSH deficiency is rarely, if ever, seen. When hypothyroidism is due to a pituitary disorder, the disease is called “secondary hypothyroidism,” and when due to a hypothalamic disorder (see p. 195) it is called “tertiary hypothyroidism.” Most patients with acquired pituitary TSH deficiency have symptoms of hypothyroidism as well as symptoms of LH, FSH, GH, and usually ACTH deficiency. Serum free T4 and T3 levels are low in association with a low or low/normal basal TSH level. The low basal TSH level fails to respond to TRH stimulation. It is often difficult to distinguish secondary and tertiary hypothyroidism. The basal levels of free T4, free T3 and TSH are similar in either etiology. However, in secondary hypothyroidism, the TSH response to TRH is usually completely absent [488,503–505] (Fig. 5.21). In contrast, tertiary hypothyroidism usually demonstrates a TSH response to TRH stimulation which is not absent, but rather delayed and attenuated, resulting in peak TSH values at 60 to 180 minutes after injection of TRH [488,506–510]. These patterns are different from that seen in normal controls and primary hypothyroidism, in which the peak TSH response to TRH stimulation is between 20 and 30 minutes follow-

20

22

0

Patients

198

FIGURE 5.23. The development of the hormonal deficiencies in 35 patients with gene mutations of the transcription factor Prophet of Pit-1 (PROP-1). The time of ocurrence of the specific hormone deficiency is indicated by the age of the patients in years. The data is expressed as a percentage of the patients on the left axis and as the number of patients on the right axis. All patients eventually showed evidence of all deficiencies, but these developed gradually over the years. GH, growth hormone; TSH, thyroid stimulating hormone; LH/FSH: luteinizing hormone/follicle stimulating hormone. From Deladoëy et al. [21].

ing the injection of TRH. Nevertheless, the TSH response to TRH is not a perfect discriminator between secondary and tertiary hypothyroidism, or between these disorders and a normal response [480,490,508]. The nocturnal surge of TSH found in normal subjects (see p. 183) does not occur in patients with either secondary or tertiary hypothyroidism, and its absence may prove to be a superior way of identifying patients with these disorders [208,491,492]. Finally, the basal serum prolactin concentration can sometimes be helpful in differentiating secondary from tertiary hypothyroidism. In secondary hypothyroidism the prolactin is usually low or normal if the destructive pituitary process is not a prolactinoma. In contrast, in tertiary hypothyroidism there is disruption of normal hypothalamic function, including the dopaminergic inhibitory tone on lactotrope cells, resulting in elevated serum prolactin levels. Increased Pituitary TSH Production–Primary

Most cases of elevated serum TSH levels are a result of primary thyroidal disease, not primary pituitary disease. However, two important causes of primary pituitary disease resulting in elevated levels of serum TSH have been described and extensively studied. TSH-secreting Pituitary Tumors

TSH-secreting pituitary tumors are rare neoplasms of the anterior pituitary. They comprise less than 1% of all pituitary tumors [511]. The clinical presentation, diagnostic work up, and therapeutic approach to TSH-secreting pituitary tumors will be discussed in Chapter 13. These tumors are characterized by a slow and indolent growth. Phenotypically, the cells are quite differentiated and synthesize the TSH b- and a-subunits characteristic of the normal thyrotropes. However, the a-subunit is synthesized in excess of the TSH b-subunit [460,462]. This phenomenon is useful in that a molar ratio of a-subunit : TSH of greater than 1 supports the diagnosis of a TSH-secreting pituitary tumor. The

Chapter 5

molecular basis for increased transcription of both the a- and TSH b-subunit genes and the excessive a-subunit gene transcription relative to the TSH b-subunit gene is

unknown. However, the defect does not appear to be the result of aberrantly initiated mRNAs for the TSH b- or asubunit. Both TSH b- and a-subunit mRNAs appear to be of sizes similar to those found in normal TSH cells [512]. Furthermore, the transcriptional initiation site of the TSH b-subunit has been mapped in a human TSH-secreting tumor and is consistent with the authentic start-site of normal TSH b-subunit gene transcription [512]. TSH-secreting tumors fail to respond to TRH stimulation [513–515] (Fig. 5.24). This reproducible finding might implicate a constitutive TRH receptor activation in the pathogenesis of these tumors. Detailed studies of the TRH receptor have not been performed in TSH-secreting tumors, but it is conceivable that there is constitutive activation of the TRH receptor due to a somatic cell mutation in the TRH receptor gene of the thyrotrope, similar to the mutation of the TSH receptor gene found in hyperfunctioning thyroid nodules [388]. Another characteristic of these tumors is their failure to respond to thyroid hormone by the normal negative feedback of thyroid hormone on TSH production, suggesting an abnormality in the TR. The altered TR expression may result from a posttranscriptional defect, since in two TSH-secreting tumors examined TRb and TRa mRNA levels were normal but the respective proteins were not detected [516]. Nevertheless, the combination of absent TRH and thyroid hormone responsiveness suggests that the defect in TSH-secreting tumors is very basic, capable of altering processes originating at the cell membrane (TRH response) and within the cell nucleus (T3 response). In contrast, inhibition of TSH release in response to SRIH is preserved in these tumors [511] (Fig. 5.24). This suggests that the pathogenetic mechanism resides proximal to the processes of hormone release. Thyroid Hormone Resistance Syndromes

Another group of pituitary gland disorders resulting in elevated levels of serum TSH are the thyroid hormone resis-

Thyroid-stimulating Hormone

199

tance syndromes [517–519]. In 1967, Refetoff et al. [520] were the first to describe three siblings who were clinically euthyroid or hypothyroid with goiters, stippled epiphyses, and deaf mutism. Each of the children had elevated levels of protein-bound iodide which were subsequently shown to be associated with high serum total and free thyroid hormone levels, elevated TSH levels, and peripheral tissue responses that were refractory to not only the endogenous high levels of thyroid hormone, but also exogenously administered supraphysiological levels of thyroid hormone [518]. Since 1967 over 700 cases of thyroid hormone resistance belonging to about 250 families have been identified [521,522]. It has become clear that there is a wide clinical spectrum in the phenotypic presentation of this disorder. At one end were cases exhibiting thyroid hormone resistance in virtually every tissue studied. In contrast, other cases were characterized more by pituitary resistance to thyroid hormone with peripheral responses similar to hyperthyroidism. Thus, the concept arose that there are two clinically distinct disorders: one, generalized resistance to thyroid hormone (GRTH) (Fig. 5.25), and the other, selective pituitary resistance to thyroid hormone (PRTH) (Fig. 5.26) [513,514]. This distinction is clinically important because cases with selective PRTH have clinical hyperthyroidism and warrant treatment of the hyperthyroidism, whereas those with GRTH are generally euthyroid and sometimes, in fact, hypothyroid. In these cases, treatment of hyperthyroxinemia with antithyroid drug therapy, radioactive iodine or surgery would be inappropriate [523]. Both disorders represent diagnostic dilemmas. Clinical presentations are often not characteristic of classic hyperthyroidism or hypothyroidism. Yet, the laboratory abnormalities are often quite spectacular, with dramatic elevations of thyroid hormone levels, inappropriately normal or elevated serum TSH levels, and elevated radioactive iodine uptake, with diffuse homogenous uptake in an enlarged thyroid gland [512–514]. The molecular basis in most cases of GRTH and PRTH is an abnormality in the thyroid hormone receptor (TR). There are two known TR genes: the TRa gene on

FIGURE 5.24. Thyroid-stimulating hormone (TSH) responses to various stimulation and suppression tests in a patient with a TSH pituitary tumor. Thyrotropinreleasing hormone (TRH) (500 mg intravenously) resulted in no TSH response, dopamine (4 mg/minute for 4 hours) resulted in no suppression of TSH secretion, SRIH (SRIF, 500 mg bolus followed by 250 mg/minute for 4 hrs) resulted in significant suppression of serum TSH levels.

200

SECTION 1

Hypothalamic–Pituitary Function

FIGURE 5.25. Generalized resistance to thyroid hormone is characterized by resistance to the action of thyroid hormone at both pituitary and peripheral tissues. The patient is euthyroid clinically with increased total and free thyroid hormone levels, and an increased or normal thyroid-stimulating hormone (TSH). From McDermott and Ridgway [517].

FIGURE 5.26. Selective pituitary resistance to thyroid hormone is characterized by resistance to the action of thyroid hormone at the level of the pituitary, but not peripheral tissues. Patients are hyperthyroid clinically, with elevated total and free thyroid hormone levels and increased or normal thyroid-stimulating hormone (TSH) levels. From McDermott and Ridgway [517].

FIGURE 5.27. Mutations of the T3 receptor b (TRb) gene in patients with generalized resistance to thyroid hormone (GRTH) are located in exons 9 and 10 which encode the T3 binding domain of the TRb receptor. From McDermott and Ridgway [517].

chromosome 17 and the TRb gene on chromosome 3 [524]. Both genes encode two different isoform products using alternative mRNA splicing. The TRa gene produces TRa1 and an a2 isoform which fails to bind thyroid hormone. The TRb gene produces two isoforms, TRb1 and TRb2 proteins, both of which bind thyroid hormone. The TRb2 isoform appears to be selectively expressed in the anterior pituitary gland and certain parts of the central nervous system. All thyroid hormone receptor isoforms are intranuclear proteins which interact with the DNA of genes influenced by thyroid hormone. Binding of thyroid hormone the receptor stimulates or suppresses transcription of the targeted gene. GRTH was found to be linked to the TRb gene locus

on chromosome 3 [525], and was then localized to point mutations in the ninth and tenth exons of the TRb gene [521,525,526] which encode for the T3-binding and adjacent hinge domains [517,527] (Fig. 5.27). These mutations usually disrupt normal T3 binding without altering DNA binding. Interaction with corepressors and coactivators has been shown to be altered in some cases [124,528,529]. In addition, GRTH has been described to occur in the absence of TRb or TRa mutations [522,530]. These cases may result from a mutation in one of those cofactors, analogous to mice deficient in steroid receptor coactivator 1, that exhibit a similar phenotype [123]. Most cases of GRTH are heterozygotes and inherited as autosomal dominant traits, with

Chapter 5

only half of their TRb receptors being abnormal. The mutated TRb1 receptor protein has been shown to be able to interfere with the function of other normal TRb1, TRb2 or TRa1 receptors, resulting in a dominant negative effect [526,531–533]. Rare individuals who are homozygous for the mutation have been reported and, in general, are more severely affected [534,535]. Cases of GRTH have also been described where a major complete TRb gene deletion results in the absence of TRb receptors. In these cases, the disorder was transmitted as an autosomal recessive and only homozygotes were affected [536]. TRa gene mutations have not been reported in GRTH [517]. The mutations in patients with PRTH have not been as widely studied. However, TRb gene mutations in the T3 binding domain have been found in PRTH [537–539]. Mutations found to be associated with PRTH appear to affect the activity of the pituitary-specific isoform TRb2 and not TRb1 [117]. However, it is interesting that patients classified as either GRTH or PRTH have been found to contain the same point mutation [537,538], and patients within a given kindred, thus having the same mutation, can also present phenotypically with either GRTH or PRTH. This phenotypic heterogeneity underlies the view that both syndromes represent different ends of the clinical spectrum of a common genetic disorder. Yet, it is helpful to separate these syndromes to direct the differential dignosis and clinical management.

GRTH In GRTH, the degree of resistance may vary from tissue to tissue, both within an individual and within a family. Affected individuals may be asymptomatic and discovered incidentally, whereas others may have nonspecific symptoms, mental retardation, short stature, hyperactivity, or even symptoms suggestive of hyperthyroidism [540–543].

Table 5.3. [517]

Thyroid-stimulating Hormone

The differential diagnosis of a clinically euthyroid subject with elevated total T4 and normal or elevated TSH can be difficult [544,545] (Table 5.3). It is also sometimes difficult to be sure whether the patient is in fact euthyroid, hypothyroid or hyperthyroid. When clinical findings are inconclusive, a variety of tissue markers may be useful. These include basal metabolic rate, cardiac systolic time intervals, serum levels of angiotensin converting enzyme, sex hormone binding globulin, ferritin, and gastrin. If all of these peripheral measures of thyroid hormone action are normal, then the patient can be categorized as having clinical euthyroidism. It is important then to exclude abnormalities of T4 binding by altered T4-binding proteins. If the patient also has no acute medical or psychiatric illness and is not taking amphetamines or amiodarone, then a presumptive diagnosis of GRTH may be made. This diagnosis can be confirmed by demonstrating that suppression of pituitary TSH secretion requires supraphysiological levels of T4 or T3. If the diagnosis of GRTH is made, treatment is generally contraindicated and may be harmful [523]. The increased thyroid hormone levels seen in this disorder are an appropriate and compensatory response to the elevated TSH levels that arise as a result of the pituitary resistance to circulating levels of thyroid hormone. Inadvertent administration of anti-thyroid drugs, radioactive iodine or surgical thyroidectomy needs to be avoided since these therapeutic modalities will lower thyroid hormone levels and result in hypothyroidism. Such iatrogenically induced hypothyroidism has clinical consequences, and requires treatment with supraphysiologic doses of thyroid hormone.

PRTH There are fewer reported cases of PRTH than GRTH and in many instances of PRTH detailed family studies have not been performed [513–515,517]. Both sporadic and familial

Causes of euthyroid hyperthyroxinemia. From McDermott and Ridgway

GRTH TBG excess FDH or TBPA excess Anti-T4 antibodies Acute medical illness Acute psychiatric Amphetamines Amiodarone, ipodate, iopanioc acid

201

Total T4

Total T3

T3 resin uptake

Free T4

Free T3

TSH

TRH Test

≠ ≠ ≠ ≠, Ø ≠ ≠ ≠ ≠

≠ ≠ N N Ø N ≠ Ø

N, ≠ Ø N N Ø, N, ≠ Ø, N N N

≠ N N N Ø – – ≠

≠ N N N ≠ – – Ø

N, ≠ N N N, ≠ N N N N, ≠

N, ≠ N N N, ≠ N Ø, N N ≠

T4, thyroxine; T3, triiodothyronine; TSH, thyroid-stimulating hormone; TRH, thyrotropin-releasing hormone; TBG, thyroxine-binding globulin; FDH, familial dysalbuminemic hyperthyroxinemia; TBPA, thyroxine-binding prealbumin; N, normal.

202

SECTION 1

Hypothalamic–Pituitary Function

cases have been reported, and the inheritance patterns of PRTH are incompletely clarified [517]. Patients with PRTH have clinical thyrotoxicosis and the evidence for clinical thyrotoxicosis is often overwhelming, including elevated levels of basal metabolic rate, fast cardiac systolic time intervals, and elevated serum concentrations of sex hormone binding globulin and angiotensin-converting enzyme. In other cases, the clinical presentation is much more subtle with only minor abnormalities found in the peripheral responses to thyroid hormone, for example, only a fast cardiac pulse rate. In the laboratory evaluation of PRTH, the TSH level is normal or elevated, a finding that clearly distinguishes it from all other primary thyroid disorders causing hyperthyroidism. Only one other disorder has clinical hyperthyroidism, high levels of thyroid hormone and inappropriately normal or elevated TSH values: the TSHsecreting tumor. Thus, a distinction between these two disorders is mandatory (Table 5.4). In TSH-secreting tumors, TSH fails to respond to TRH stimulation, whereas in PRTH there is an exuberant response [513–515]. Furthermore, patients with TSH-secreting tumors have elevated concentrations of free a-subunit with a-subunit : TSH molar ratios of greater than 1, whereas patients with PRTH have a-subunit : TSH molar ratios less than or equal to 1 [460–462,511]. Finally, patients with TSH-secreting pituitary tumors most commonly have very abnormal pituitary glands morphologically reflected in abnormal structural studies of the pituitary, whereas cases with PRTH have normal pituitary radiographic images [511]. Since patients with PRTH and TSH-secreting pituitary tumors both have clinical hyperthyroidism, treatment is warranted. Direct ablation of the thyroid gland by radioactive iodine or surgery has been used successfully in both. Ideally, TSH-secreting pituitary tumors should be treated first with transsphenoidal resection in an attempt to remove the tumorous source of excessive TSH production. In contrast, pituitary surgery for PRTH would not be successful unless a total hypophysectomy was performed. Since TSH overproduction is the primary defect in PRTH, many attempts

Table 5.4. Causes of thyroid-stimulating hormone (TSH)mediated hyperthyroidism. From McDermott and Ridgway [517]

Symptoms Total and free T4 Total and free T3 TSH TSH response to TRH a-Subunit a/TSH molar ratio

PRTH

TSH Tumor

Thyrotoxicosis ≠ ≠ N/≠ ≠ N £1

Thyrotoxicosis ≠ ≠ N/≠ Flat ≠/N >1

T4, thyroxine; T3, triiodothyronine; TRH, thyrotropin-releasing hormone; PRTH, pituitary resistance to thyroid hormone; a/TSH molar ratio, (a-subunit [ng/ml] ∏ TSH [mU/l] ¥ 10); N, normal.

have been made to develop alternative medical therapies to suppress the TSH hypersecretion. These include L-T3 [546], D-T4 [547,548], and triiodothyrocetic acid administration [549–551]. None of these have been universally successful. More recently, the use of octreotide, a SRIH analog, and bromocriptine have both been shown to acutely reduce TSH and thyroid hormone levels [552–558]. Unfortunately, neither have been effective over longer therapeutic intervals and rebound elevations of the serum thyroid hormone levels, with escape from pituitary TSH suppression, have occurred. Altered Pituitary TSH Production–Secondary

The most common way that pituitary TSH production is influenced is via its inverse relationship with circulating levels of thyroid hormone. Thus, elevated levels of free T4 and free T3 cause a decrease in pituitary TSH synthesis and secretion. In contrast, decreases in circulating free T4 or free T3 levels cause a dramatic augmentation in pituitary TSH production (see pp. 178 and 187). In human studies, measurements of serum TSH levels using immunometric assays have shown an inverse linear relationship between serum TSH levels and circulating levels of thyroid hormone (Fig. 5.19). Patients with clinical hypothyroidism have dramatic elevations of serum TSH, whereas those with clinical hyperthyroidism have low or absent levels of circulating TSH. As the measurements for TSH have improved from first to fourth generation TSH assays, patients with hyperthyroidism have been found to have lower and lower serum TSH levels [449–452] (Fig. 5.18). Using fourth generation TSH assays, patients with hyperthyoidism either have TSH levels below the detection level of 0.001 mU/l, or very low levels in the range of 0.001 to 0.005 mU/l [451]. These extraordinarily low serum TSH levels found in hyperthyroidism suggest that the thyrotrope is exquisitely sensitive to elevated levels of thyroid hormone. The mechanisms responsible for this are currently under active investigation. Thyroid hormone decreases transcription from the TRH gene in the hypothalamus [297,298], and at the level of the pituitary it decreases both the a and TSHb gene transcription [101,102] (see p. 178). In hypothyroidism, transcription of the a- and TSHb-subunit genes has been found to be dramatically elevated [101,102,112,559]. Thyroid hormone also inhibits TSH secretion from the thyrotrope (see p. 187). In fact, it transiently increases cellular content of TSH [111,560]. Thyroid hormone influences TSH secretory pulse amplitude but not frequency [305,561]. Finally, elevated levels of thyroid hormone cause a 20% to 30% increase in TSH metabolic clearance rate, whereas the opposite is seen with hormone deficiency [188]. Thus, the effects of thyroid hormone on TSH serum profiles are multifactorial, involving effects at the level of the hypothalamus, pituitary and peripheral TSH clearance. The differential diagnosis of a low serum TSH level includes any primary thyroid disorder causing elevated

Chapter 5

circulating levels of thyroid hormone (Table 5.5). The most common cause is excessive thyroid hormone administration. Low serum TSH levels may be found in euthyroid sick syndrome during the early acute phases, in various causes of hyperthyroidism, or following the administration of superphysiological dopamine or glucocorticoids, as well as hypopituitarism. In contrast, an elevated serum TSH level may be due to any primary thyroid disorder which lowers circulating levels of thyroid hormone (Table 5.6). In the USA, the most common causes are autoimmune thyroiditis, ablation of the thyroid gland by radioactive iodine, surgical removal of the thyroid gland, or antithyroid drugs. In third world countries, iodine deficiency is still a common cause of defective thyroid hormone production and elevated serum TSH levels. Rarely, it is due to a TSH-secreting pituitary tumor. Subclinical Hyperthyroidism and Hypothyroidism

Subclinical hyperthyroidism and subclinical hypothyroidism are terms that refer to clinical syndromes in which the only abnormality in the diagnostic evaluation is an alteration of the serum TSH concentration. Specifically, in subclinical hyperthyroidism the TSH level is low, usually ranging between 0.005 and 0.5 mU/l, whereas in subclinical hypothyroidism the basal serum TSH concentration is elevated, usually greater than 5 mU/l. The patients generally do not present with clinical symptoms that would be diagnostic of either hyperthyroidism or hypothyroidism. Further-

Table 5.5. Differential diagnosis of a low sensitive thyroidstimulating hormone level L-thyroxine therapy Euthyroid sick syndrome Hyperthyroidism Graves’ disease Hyperfunctioning thyroid nodule Hyperfunctioning multinodular goiter Subacute thyroiditis Painless thyroiditis Glucocorticoids Dopamine Hypopituitarism

Table 5.6. Differential diagnosis of a high sensitive thyroidstimulating hormone (TSH) level Autoimmune thyroiditis Ablation of the thyroid gland Radioactive iodine Thyroidectomy Anti-thyroid drugs Iodine deficiency TSH-secreting pituitary tumors

Thyroid-stimulating Hormone

203

more, the serum concentrations of free T4 and free T3 are within the normal range. Thus, the subclinical designation refers to the fact that physicians cannot make a clinical diagnosis because the disorder is either very mild or early in its evolution. These disorders have been widely identified because the immunometric TSH assays amplify the sensitivity and specificity of abnormal serum TSH levels. The sensitivity of the assays is so highly developed that abnormalities in TSH secretion are found before serum circulating thyroid hormone levels fall or rise outside of the normal range, and certainly before classical clinical symptoms occur. The differential diagnosis of subclinical hyperthyroidism and hypothyroidism is the same as that described above for the clinical disorders (Tables 5.5 and 5.6). However, the most common cause of subclinical hyperthyroidism is inadvertent administration of excessive amounts of thyroid hormone. Iatrogenic subclinical hyperthyroidism is very common and may occur in as many as a third of patients taking thyroid hormone for replacement therapy [450]. More importantly, subclinical hyperthyroidism has long-term consequences and is not merely a biochemical curiosity. Premature osteoporosis, increased cardiac chronotropy and ionotropy, increased cardiac atrial premature beats, increased left ventricular mass and deteriorating quality of life scores all provide compelling evidence that a low serum TSH level represents an important biologic signal [562–564]. Conversely, the most common cause of subclinical hypothyroidism is inadequate treatment of primary hypothyroidism with thyroid hormone, that occurs in as many as a fifth of patients on thyroid hormone replacement therapy [450]. The abnormal serum TSH level in subclinical hypothyroidism reflects important biological disturbances including reduced cardiac chronotropy and ionotropy, increased serum lipid levels, decreased neurophysiological performance, and decreased quality of life [565–569]. Recently, Haddow et al. reported that mild hypothyroidism in pregnant women (mean TSH 13 mU/l) was associated with slightly lower neurophysiological performance in the euthyroid offspring evaluated through age 7 to 9, and showed that treatment, even when inadequate, resulted in an improvement [570]. This suggests that correcting subclinical hypothyroidism may be critical for the early stages of development. In conclusion, controversy still exists on the question of whether isolated abnormalities in serum TSH levels without definitive clinical symptoms warrant therapy or are merely insignificant biochemical aberrations. However, the majority of evidence would favor the view that isolated increases or decreases in serum TSH levels deserve a thorough evaluation and, in most instances, the abnormal TSH level is an important signal and needs therapeutic intervention. USE OF RECOMBINANT THYROTROPIN Using recombinant DNA technology, human TSH has been produced in Chinese hamster ovary cells and purified for

204

SECTION 1

Hypothalamic–Pituitary Function

use in human subjects [156,571]. The most important current approved indication for the use of recombinant human TSH (rhTSH) is for the diagnostic follow-up evaluation of patients who have been previously treated for differentiated thyroid cancer [572]. Two Phase III studies [573,574] have shown that daily intramuscular injections (0.9 mg) of rh TSH for 2 days produces elevated serum TSH concentrations (50–150 mU/l) which remain above 25 mU/l for 4 days. Such substantial increases in serum TSH levels provide an effective stimulus for radioactive iodine incorporation and thyroglobulin release by residual normal or malignant thyroid tissue. Current practice recommends performing radioactive iodine whole body scans and thyroglobulin measurements in the hypothyroid state when endogenous high serum TSH levels stimulate potential residual or cancerous thyroid cells. Comparisons of whole body scans and serum thyroglobulin levels during stimulation with rhTSH while the patient is taking thyroid hormone suppressive therapy versus during hypothyroidism after thyroid hormone withdrawal shows equivalent, though not identical, results [574]. The overall detection of residual normal thyroid tissue or thyroid cancer using rhTSH was 90% of that detected during hypothyroidism following thyroid hormone withdrawal. In addition, the detection of metastatic disease using rhTSH was 100% of that detected by hypothyroid whole body scanning and serum thyroglobulin measurements. Such comparative studies convincingly demonstrated that patients strongly preferred having the whole body scans and serum thyroglobulin measurements performed after rhTSH while taking thyroid hormone therapy because of the absence of hypothyroid symptomatology, resulting in an improved quality of life. Based on this evidence, rhTSH has been approved for the diagnostic follow-up evaluations of patients with treated differentiated thyroid cancer. Future uses of this product await appropriate prospective studies but may include therapy of thyroid cancer, the diagnostic evaluation of central hypothyroidism, or the therapy of cases of hyperthyroidism where the uptake may be enhanced by stimulation with rhTSH. ACKNOWLEDGMENT The authors would like to thank Linda Trefry for excellent secretarial assistance. REFERENCES 1 Voss JW, Rosenfeld MG. Anterior pituitary development: short tales from dwarf mice. Cell 1992;70:527–530. 2 Treier M, Gleiberman AS, O’Connell SM et al. Multistep signaling requirements for pituitary organogenesis in vivo. Genes and Dev 1998;12: 1691–1704. 3 Hermesz E, Mackem S, Mahon KA. Rpx: a novel anterior-restricted homeobox gene progressively activated in the prechordal plate, anterior neural plate and Rathke’s pouch of the mouse embryo. Development 1996;122:41–52. 4 Lamonerie T, Tremblay JJ, Lanctot C et al. Ptx1, a bicoid-related homeo box transcription factor involved in transcription of the pro-opiomelanocortin gene. Genes and Dev 1996;10:1284–1295.

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autoimmune thyroid disease and nonhomologous with gonadotropin receptors. Relationship of functional and immunogenic domains. J Biol Chem 1991;266:19413–19418. Stein SA, Oates EL, Hall CR et al. Identification of a point mutation in the thyrotropin receptor of the hyt/hyt hypothyroid mouse. Mol Endocrinol 1994;8:129–138. Refetoff S, Sunthorntepvarakul T, Gottschalk ME, Hayashi Y. Resistance to thyrotropin and other abnormalities of the thyrotropin receptor. Recent Prog Horm Res 1996;51:97–120. Abramowicz MJ, Duprez L, Parma J et al. Familial congenital hypothyroidism due to inactivating mutation of the thyrotropin receptor causing profound hypoplasia of the thyroid gland. J Clin Invest 1997;99: 3018–3024. Costagliola S, Sunthorntepvarakul T, Migeotte I et al. Structure-function relationships of two loss-of-function mutations of the thyrotropin receptor gene. Thyroid 1999;9:995–1000. LaFranchi S. Congenital hypothyroidism: etiologies, diagnosis, and management. Thyroid 1999;9:735–740. Grossmann M, Szkudlinski MW, Wong R et al. Substitution of the seat-belt region of the thyrotropin (TSH)-b subunit with the corresponding regions of the choriogonadotropin or follitropin confers luteotropic, but not follitropic, activity to chimeric TSH. J Biol Chem 1997;272:15532– 15540. Leinung MC, Bergert ER, McCormick DJ, Morris JC. Synthetic analogs of the carboxyl-terminus of b-thyrotropin: the importance of basic amino acids in receptor binding activity. Biochemistry 1992;31:10094–10098. Medeiros-Neto G, Herodotou DT, Rajan S et al. A circulating biologically inactive thyrotropin caused by a mutation in the b subunit gene. J Clin Invest 1996;97:1250–1256. Biebermann H, Liesenkotter KP, Emeis M et al. Severe congenital hypothyroidism due to a homozygous mutation of the bTSH gene. Pediatr Res 1999;46:170–173. Leinung MC, Reed DK, McCormick DJ et al. Further characterization of the receptor-binding region of the thyroid-stimulating hormone a-subunit. Proc Natl Acad Sci USA 1991;88:9707–9711. Grossmann M, Szkudlinski MW, Tropea JE et al. Expression of human thyrotropin in cell lines with different glycosylation patterns combined with mutagenesis of specific glycosylation sites: characterization of a novel role for the oligosaccharides in the in vitro and in vivo bioactivity. J Biol Chem 1995;270:29378–29385. Szkudlinski MW, Thotakura NR, Weintraub BD. Subunit-specific functions of N-linked oligosaccharides in human thyrotropin: role of terminal residues of a- and b-subunit in metabolic clearance and bioactivity. Proc Natl Acad Sci USA 1995;92:9062–9066. Thotakura NR, Desai RK, Szkudlinski MW, Weintraub BD. The role of the oligosaccharide chains of thyrotropin a- and b-subunits in hormone action. Endocrinology 1992;131:82–88. Renwick A, Wiggin P. An antipodean perception of the mode of action of glycoprotein hormones. FEBS Lett 1992;297:1–3. Davies TF, Platzer M. hCG-induced receptor activation and growth acceleration in FRTL-5 thryoid cells. Endocrinology 1986;118:2149–2151. Tomer Y, Huber GK, Davies TF. Human chorionic gonadotropin (hCG) interacts directly with recombinant human TSH receptors. J Clin Endo and Metab 1992;74:1477–1479. Yoshimura M, Hershman JM, Pang XP et al. Activation of the thyrotropin (TSH) receptor by human chorionic gonadotropin and luteinizing hormone in Chinese hamster ovary cells expressing functional human TSH receptors. J Clin Endo and Metab 1993;77:1009–1013. Yoshimura M, Hershman JM. Thyrotropic action of human chorionic gonadotropin. Thyroid 1995;5:425–434. Rodien P, Bremont C, Sanson ML et al. Familial gestational hyperthyroidism caused by a mutant thyrotropin receptor hypersensitive to human chorionic gonadotropin. N Engl J Med 1998;339:1823–1826. Chazenbalk GD, Nagayama Y, Russo D et al. Functional analysis of the cytoplasmic domains of the human thyrotropin receptor by site-directed mutagenesis. J Biol Chem 1990;265;20970–20975. Parmentier M, Libert F, Maenhaut C et al. Molecular cloning of the thyrotropin receptor. Science 1989;246:1620–1622. Wolff J, Jones AB. The purification of bovine thyroid plasma membranes and the properties of membrane-bound adenyl cyclase. J Biol Chem 1971;246: 3939–3947. Yamashita K, Field JB. Preparation of thyroid plasma membranes containing TSH-responsive adenyl cyclase. Biochem Biophys Res Commun 1970;40: 171–178.

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381 Philip NJ, Grollman EF. Thyrotropin and norepinephrine stimulate the metabolism of phosphoinositides in FRTL-5 thyroid cells. FEBS Lett 1986;202:193–196. 382 Field JB, Ealey PA, Marshall NJ, Cockcroft S. Thyroid-stimulating hormone stimulates increases in inositol phosphates as well as cyclic AMP in the FRTL5 rat thyroid cell line. Biochem J 1987;247:519–524. 383 Laurent E, Mockel J, Van Sande J et al. Dual activation by thyrotropin of the phospholipase C and cyclic AMP cascades in human thyroid. Mol Cell Endocrinol 1987;52:273–278. 384 Van Sande J, Parma J, Tonacchera M et al. Somatic and germline mutations of the TSH receptor gene in thyroid diseases. J Clin Endo and Metab 1995;80: 2577–2585. 385 Chazenbalk GD, Kakinuma A, Jaume JC et al. Evidence of negative cooperativity among human thyrotropin receptors overexpressed in mammalian cells. Endocrinology 1996;137:4586–4591. 386 Esapa CT, Duprez L, Ludgate M et al. A novel thyrotropin receptor mutation in an infant with severe thyrotoxicosis. Thyroid 1999;9:1005–1010. 387 Nogueira CR, Kopp P, Arseven OK et al. Thyrotropin receptor mutations in hyperfunctioning thyroid adenomas from Brazil. Thyroid 1999;9:1063– 1068. 388 Parma J, Duprez L, Van Sande J et al. Somatic mutations in the thyrotropin receptor gene cause hyperfunctioning thyroid adenomas. Nature 1993; 365:649–651. 389 Gabriel EM, Bergert ER, Grant CS et al. Germline polymorphism of codon 727 of human thyroid-stimulating hormone receptor is associated with toxic multinodular goiter. J Clin Endo and Metab 1999;84:3328– 3335. 390 Kosugi S, Okajima F, Ban T et al. Mutation of alanine 623 in the third cytoplasmic loop of the rat thyrotropin (TSH) receptor results in a loss in the phosphoinositide but not cAMP signal induced by TSH and receptor autoantibodies. J Biol Chem 1992;267:24153–24156. 391 Cooper E, Spaulding SW. Hormonal control of the phosphorylation of histones, HMG protein and other nuclear proteins. Mol Cell Endocrinol 1985;39:1–20. 392 Contor L, Lamy F, Lecocq R et al. Differential protein phosphorylation in induction of thyroid cell proliferation by thyrotropin, epidermal growth factor or phorbol ester. Mol Cell Biol 1988;8:2494–2503. 393 Ericson LE. Exocytosis and endocytosis in the thyroid follicle cell. Mol Cell Endocrinol 1981;22:1–24. 394 Nilsson M, Engstrom G, Ericson LE. Graded response in the individual thyroid follicle cell to increasing doses of TSH. Mol Cell Endocrinol 1986;44:165–169. 395 Collins WT, Capen CC. Ultrastructural and functional alterations of the rat thyroid gland produced by polychlorinated biphenyls compared with iodide excess and deficiency, and thyrotropin and thyroxine administration. Virchows Arch 1980;33:213–231. 396 Nielsen TB, Ferdows MS, Brinkley BR, Field JB. Morphological and biochemical responses of cultured thyroid cells to thyrotropin. Endocrinology 1985;116:788–797. 397 Roger PP, Rukaert F, Lamy F et al. Actin stress fiber disruption and tropomyosin isoform switching in normal thyroid epithelial cells stimulated by thyrotropin and phorbol esters. Exp Cell Res 1989;182:1–13. 398 Knopp J, Stolc V, Tong W. Evidence for the induction of iodide transport in bovine thyroid cells treated with thyroid stimulating hormone or dibutyryl cyclic adenosine 3¢5¢ monophosphate. J Biol Chem 1970;245:4403–4408. 399 Damante G, Chazenbalk G, Russo D et al. Thyrotropin regulation of thyroid peroxidase mesenger ribonucleic acid levels in cultured rat thyroid cells: evidence for involvement of a non-transcriptional mechanism. Endocrinology 1989;124:2889–2894. 400 Corda D, Kohn LD. Phorbol myristate acetate inhibits adrenergically but not thyrotropin-regulated function in FRTL-5 rat thyroid cells. Endocrinology 1987;120:1152–1160. 401 Burke G. Effects of iodide on thyroid stimulation. J Clin Endo and Metab 1970;30:76–84. 402 Tosta Z, Chabaud O, Chebath J. Identification of thyroglobulin mRNA sequences in the nucleus and cytoplasm of cultured thyroid cells: a fast transcriptional effect of thyrotropin. Biochem Biophys Res Commun 1983; 116:54–61. 403 Bybee A, Tuffery AR. Rapid proliferative response of rat thyroid gland to a single injection of TSH in vivo. J Endocrinol 1989;121:27–30. 404 Ealey P, Ahene CA, Emmerson JM, Marshall NJ. Forskolin and thyrotropin stimulation of rat FRTL-5 thyroid cell growth: the role of cyclic AMP. J Endocrinol 1987;114;199–205. 405 Van Sande J, Lefort A, Beebe S. Pairs of cyclic AMP analogues that are

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specifically synergistic for type I and type II cAMP dependent protein kinase mimic thyrotropin effects on the function, differentiation expression and mitogenesis of dog thyroid cells. Eur J Biochem 1989;183:699–708. Karsenty G, Alquier C, Jelsema C, Weintraub BD. Thyrotropin induces growth and iodothyronine production in a human thyroid cell line without affecting adenosine 3¢5¢ monophosphate production. Endocrinology 1988;123: 1977–1983. Tramontano D, Moses AC, Veneziani BM, Ingbar SH. Adenosine 3¢5¢monophosphate mediates both the mitogenic effect of thyrotropin and its ability to amplify the response to insulin-like growth factor I in FRTL-5 cells. Endocrinology 1988;122:127–132. Westermark K, Westermark B, Karlsson FA, Ericson LE. Location of epidermal growth factor receptors in porcine thyroid follicle cells and receptor regulation by thyrotropin. Endocrinology 1986;118:1040–1046. Tramontano D, Chin WW, Moses AC, Ingbar SH. Thyrotropin and dibutyryl cyclic AMP increase levels of c-myc and c-fos mRNAs in cultured rat thyroid cells. J Biol Chem 1986;261:3919–3922. Kawakami A, Eguchi K, Matsouka N et al. Thyroid-stimulating hormone inhibits Fas antigen-mediated apoptosis of human thyrocytes in vitro. Endocrinology 1996;137:3163–3169. Tilly JL, Tilly KI, Kenton ML, Johnson AL. Expression of members of the Bcl-2 gene family in the immature rat ovary: equine gonadotropin-mediated inhibition of granulosa cell apoptosis is associated with decreased Bax and constitutive Bcl-2 and Bcl-xlong messenger ribonucleic acid levels. Endocrinology 1995;136:232–241. Tilly KI, Banerjee S, Banerjee PP, Tilly JL. Expression of the p53 and Wilms’ tumor suppressor genes in the rat ovary: gonadotropin repression in vivo and immunohistochemical localization of nuclear p53 protein to apoptotic granulosa cells of atretic follicles. Endocrinology 1995;136:1394–1402. Grossmann M, Szkudlinski MW, Dias JA et al. Site-directed mutagenesis of amino acids 33–44 of the common a-subunit reveals different structural requirements for heterodimer expression among the glycoprotein hormones and suggests that cAMP production and growth promotion are potentially dissociable functions of hTSH. Mol Endocrinol 1996;10:769–779. Field JB, Dekker A, Titus G et al. In vitro and in vivo refractoriness to thyrotropin stimulation of iodine organification and thyroid hormone secretion. J Clin Invest 1979;64:265–271. Shuman SJ, Zor U, Chayoth R, Field JB. Exposure of thyroid slices to thyroid-stimulating hormone induces refractoriness of the cyclic AMP system to subsequent hormone stimulation. J Clin Invest 1976;57:1132–1141. Chazenbalk GD, Nagayama Y, Kaufman KD, Rapoport B. The functional expression of recombinant human thyrotropin receptors in non-thyroidal eukaryotic cells provides evidence that homologous desensitization to thyrotropin stimulation requires a cell-specific factor. Endocrinology 1990;127:1240–1244. Barnes NDH AB, Ryan RJ. Sexual maturation in juvenile hypothyroidism. Mayo Clin Proc 1973;48:849–856. Anasti JN, Flack MR, Froelich J et al. A potential novel mechanism for precocious puberty in juvenile hypothyroidism. J Clin Endo and Metab 1995;80:276–279. Nagayama Y, Yamasaki H, Takeshita A et al. Thyrotropin binding specificity for the thyrotropin receptor. J Endocrinol Invest 1995;18:283–287. Paschke R, Vassart G, Ludgate M. Current evidence for and against TSH receptor being the common antigen in Graves’ disease and thyroid associated ophthalmopathy. Clin Endocrinol (Oxf.) 1995;42:565–569. Peele ME, Carr FE, Baker JR et al. TSHb subunit gene expression in human lymphocytes. Am J Med Sci 1993;305:1–7. Endo T, Ohta K, Haraguchi K, Onaya T. Cloning and functional expression of thyrotropin receptor cDNA from fat cells. J Biol Chem 1995;270: 10833–10837. Haraguchi K, Shimura H, Kawaguchi A et al. Effects of thyrotropin on the proliferation and differentiation of cultured rat preadipocytes. Thyroid 1999;9:613–619. Morris JC. Structure and function of the TSH receptor: its suitability as a target for radiotherapy. Thyroid 1997;7:253–258. Weintraub BD, Szkudlinski MW. Development and in vitro characterization of human recombinant thyrotropin. Thyroid 1999;9:447–450. McKenzie JM. The bioassay of thyrotropin in serum. Endocrinology 1958; 63:372–382. Bakke JL, Lawrence N, Arnett F, MacFadden W. The fractionation of exogenous and endogenous thyroid-stimulating hormone from human and rat plasma and tissues. J Clin Endo and Metab 1961;21:1280–1289. D’Angelo SA. Blood thyrotropin levels in thyrotoxic patients before and after hypophysectomy. J Clin Endo and Metab 1963;23:229–234.

Chapter 5 429 Utiger RD, Odell WD, Condliffe PG. Immunologic studies of purified human and bovine thyrotropin. Endocrinology 1963;73:359–365. 430 Odell WD, Wilber JF, Paul WE. Radioimmunoassay of thyrotropin in human serum. J Clin Endo and Metab 1965;25:1179–1188. 431 Utiger RD. Radioimmunoassay of human plasma thyrotropin. J Clin Invest 1965;44:1277–1286. 432 Odell WD, Wilber JF, Utiger RD. Studies of thyrotropin physiology by means of radioimmunoassay. Recent Prog Horm Res 1967;23;47–78. 433 Mayberry WE, Gharib H, Bilstad JM. Radioimmunoassay for human thyrotropin: clinical value in patients with normal and abnormal thyroid function. Ann Int Med 1971;74:471–480. 434 Hershman JM, Pittman Jr JA. Utility of the radioimmunoassay of serum thyrotropin in man. Ann Int Med 1971;74:481–490. 435 Patel YC, Burger HG, Hudson B. Radioimmunoassay of serum thyrotropin: sensitivity and specificity. J Clin Endo and Metab 1971;33:768–774. 436 Ridgway EC, Weintraub BD, Cevallos JL et al. Suppression of pituitary TSH secretion in the patient with a hyperfunctioning thyroid nodule. J Clin Invest 1973;52:2783–2792. 437 Nisula BC, Louvet J-P. Radioimmunoassay of thyrotropin concentrated from serum. J Clin Endo and Metab 1978;46:729–733. 438 Wehmann RE, Rubenstein HA, Nisula BC. A sensitive, convenient radioimmunoassay procedure which demonstrates that serum hTSH is suppressed below the normal range in thyrotoxic patients. Endocr Res Commun 1979;6:249–255. 439 Wehmann RE, Gregerman RI, Burns WH et al. Suppression of thyrotropin in the low-thyroxine state of severe nonthyroidal illness. N Engl J Med 1985; 312:546–552. 440 Ridgway EC, Ardisson LJ, Meskell MJ et al. Monoclonal antibody to human thyrotropin. J Clin Endo and Metab 1982;55:44–48. 441 Ridgway EC. Thyrotropin radioimmunoassays: Birth, life and demise. Mayo Clin Proc 1988;63(10):1028–1034. 442 Ishikawa E, Imagawa M, Yoshitake S et al. Major factors limiting sensitivity of sandwich enzyme immunoassay for ferritin, immunoglobulin E and thyroidstimulating hormone. Ann Clin Biochem 1982;19:379–384. 443 Seth J, Kellett HA, Caldwell G et al. A sensitive immunoradiometric assay for serum thyroid stimulating hormone: a replacement for the thyrotrophin releasing hormone text? Br Med J 1984;289:1334–1336. 444 Wood WG, Waller D, Hantke U. An evaluation of six solid-phase thyrotropin (TSH) kits. J Clin Chem Clin Biochem 1985;23:461–471. 445 Odell WD, Griffin J, Zahradnik R. Two-monoclonal-antibody sandwich-type assay for thyrotropin, with use of an avidin-biotin separation technique. Clin Chem 1986;32:1873–1878. 446 Van Heyningen V, Abbott SR, Daniel SG et al. Development and utility of a monoclonal-antibody-based, highly sensitive immunoradiometric assay of thyrotropin. Clin Chem 1987;33:1387–1390. 447 Klee GG, Hay ID. Assessment of sensitive thyrotropin assays for an expanded role in thyroid function testing proposed criteria for analytic performance and clinical utility. J Clin Endo and Metab 1987;64:461–471. 448 Ross DS. New sensitive immunoradiometric assays for thyrotropin. Ann Int Med 1986;104:718–720. 449 Spencer CA, LoPresti JS, Patel A et al. Applications of a new chemiluminometric thyrotropin assay to subnormal measurement. J Clin Endo and Metab 1990;70:453–460. 450 Ross DS, Daniels GH, Gouveia D. The use and limitations of a chemiluminescent thyrotropin assay as a single thyroid function test in an outpatient endocrine clinic. J Clin Endo and Metab 1990;71:764–769. 451 Spencer CA, Schwarzbein D, Guttler RB et al. Thyrotropin (TSH)-releasing hormone stimulation test responses employing third and fourth generation TSH assays. J Clin Endo and Metab 1993;76:494–498. 452 Spencer C. New Roles for TSH Measurement in Thyroid Testing. Thyroideducational Series, Monograph 4, Rochester: Kodak Clinical Diagnostics Ltd., 1992. 453 Kourides IA, Weintraub BD, Levko MA et al. A and b subunits of human thyrotropin: purification and development of specific radioimmunoassays. Endocrinology 1974;94:1411–1421. 454 Kourides IA,Weintraub BD, Ridgway EC, Maloof F. Increase in the b subunit of human TSH in hypothyroid serum after thyrotropin releasing hormone. J Clin Endo and Metab 1973;37:836–840. 455 Kourides IA, Weintraub BD, Ridgway EC, Maloof F. Pituitary secretion of free a and b-subunit of human thyrotropin in patients with thyroid disorders. J Clin Endo and Metab 1975;40:872–885. 456 Bigos ST, Ridgway EC, Kourides IA, Maloof F. The spectrum of pituitary alterations with mild and severe thyroid impairment. J Clin Endo and Metab 1978;46:317–325.

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457 Kourides IA, Weintraub BD, Re RN et al. Thyroid hormones, estrogen, and glucocorticoid effect on two different pituitary glycoprotein hormone asubunit pools. Clin Endocrinol (Oxf.) 1978;9:535–542. 458 Kourides IA, Ridgway EC, Maloof F. Discordant responses of TSH and its free a and b-subunits after TRH with incremental thyroid hormone replacement in primary hypothyroidism. J Clin Endo and Metab 1979;49:700–705. 459 Grover AS, Griffin J, Odell W. An ultra-sensitive, specific, 2nd antibody immunoradiometric assay for the free a subunit. Clin Chem 1986;32(10):1873–1878. 460 Kourides IA, Weintraub BD, Rosen SW et al. Secretion of a-subunit of glycoprotein hormones by pituitary adenomas. J Clin Endo and Metab 1976;43:97–106. 461 Kourides IA, Re RN, Weintraub BD et al. Metabolic clearance and secretion rates of subunits of human thyrotropin. J Clin Invest 1977;59:508– 516. 462 Kourides IA, Ridgway EC, Weintraub BD et al. Thyrotropin-induced hyperthyroidism: use of a and b-subunit levels to identify patients with pituitary tumors. J Clin Endo and Metab 1977;45:534–543. 463 Ridgway EC, Klibanski A, Ladenson PW et al. Pure a-secreting pituitary adenomas. N Engl J Med 1981;304:1254–1259. 464 Klibanski A, Ridgway EC, Zervas NT. Pure a-subunit secreting pituitary tumors. J Neurosurg 1983;59:585–589. 465 Borges JLC, Ridgway EC, Kovacs K et al. Follicle-stimulating hormonesecreting pituitary tumor with concomitant elevation of serum a-subunit levels. J Clin Endo and Metab 1984;58:937–941. 466 Vance ML, Ridgway EC, Thorner MO. FSH and a-subunit secreting pituitary tumor treated with bromocriptine. J Clin Endo and Metab 1985;61:580–584. 467 Klibanski A, Deutsch PJ, Jameson JL et al. Luteinizing hormone-secreting pituitary tumor: biosynthetic characterization and clinical studies. J Clin Endo and Metab 1987;64:536–542. 468 Blackman MR, Weintraub BD, Rosen SW et al. Human placental and pituitary glycoprotein hormones and their subunits as tumor markers: a quantitative assessment. J Natl Cancer Inst 1980;65:81–93. 469 Muggia FM, Rosen SW, Weintraub BD et al. Ectopic placental proteins in nontrophoblastic tumors: serial measurements following chemotherapy. Cancer 1975;36:1327–1337. 470 Kahn CR, Rosen SW, Weintraub BD et al. Ectopic production of chorionic gonadotropin and its subunits by islet cell tumors: a specific marker for malignancy. N Engl J Med 1977;197:565–569. 471 Bender RA, Weintraub BD, Rosen SW. Prospective evaluation of two tumorassociated proteins in pancreatic adenocarcinoma. Cancer 1979;45:591–595. 472 Broder LE, Weintraub BD, Rosen SW et al. Placental proteins and their subunits as tumor markers in prostatic carcinoma. Cancer 1977;40:211–216. 473 Metz SA, Weintraub BD, Rosen SW et al. Ectopic secretion of chorionic gonadotropin by a lung carcinoma. Pituitary gonadotropin and subunit secretion and prolonged chemotherapeutic remission. Am J Med 1978; 65:325–333. 474 Tashjian Jr AH, Weintraub BD, Barowsky NJ et al. Subunits of human chorionic gonadotropin: unbalanced synthesis and secretion by clonal cell strains derived from a bronchogenic carcinoma. Proc Natl Acad Sci USA 1973;70:1419–1422. 475 Rosen SW, Weintraub BD, Aaronson SA. Nonrandom ectopic protein production by malignant cells: direct evidence in vitro. J Clin Endo and Metab 1980;50:834–841. 476 Blackman MR, Weintraub BD, Rosen SW et al. Comparison of the effects of lung cancer, benign lung disease, and normal aging on pituitary–gonadal function in men. J Clin Endo and Metab 1988;66:88–95. 477 Faglia G. The clinical impact of the thyrotropin-releasing hormone text. Thyroid 1998;8:903–908. 478 Bowers CY, Schally AV, Schalch AS et al. Activity and specificity of synthetic thyrotropin-releasing hormone in man. Biochem Biophys Res Commun 1970;39: 352–355. 479 Hershman JM, Pittman Jr JA. Response to synthetic thyrotropin-releasing hormone in man. J Clin Endo and Metab 1970;31:457–460. 480 Patel YC, Burger HG. Serum thyrotropin (TSH) in pituitary and/or hypothalamic hypothyroidism: normal or elevated basal levels and paradoxical responses to thyrotropin-releasing hormone. J Clin Endo and Metab 1973;37: 190–196. 481 Faglia G, Beck-Peccoz P, Ambrosi B et al. Prolonged and exaggerated elevations in plasma thyrotropin (TSH) after thyrotropin releasing factor (TRF) in patients with pituitary tumors. J Clin Endo and Metab 1971;33:999–1002. 482 Jackson IMD, Reichlins S. Thyrotropin-releasing hormone (TRH): distribution in hypothalamic and extrahypothalamic brain tissues of mammalian and submammalian chordates. Endocrinology 1974;95:854–862.

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483 Mallik TK, Wilber JF, Pegues J. Measurements of thyrotropin-releasing hormone like material in human peripheral blood by affinity chromatography and radioimmunoassay. J Clin Endo and Metab 1982;54:1194–1198. 484 Faglia G, Beck-Peccoz P, Ballabio M, Nava C. Excess of b-subunit of thyrotropin in patients with idiopathic central hypothyroidism due to the secretion of TSH with reduced biological activity. J Clin Endo and Metab 1983;56:908–914. 485 Beck-Peccoz P, Amr S, Menezes-Ferreira NM et al. Decreased receptor binding of biologically inactive thyrotropin in central hypothyroidism: effect of treatment with thyrotropin-releasing hormone. N Engl J Med 1985;312: 1085–1090. 486 Porter BA, Refetoff S, Rosenfield RL et al. Abnormal thyroxine metabolism in hyposomatotrophic dwarfism and inhibition of responsiveness to TRH during GH therapy. Pediatrics 1973;51:668–674. 487 Faglia G, Beck-Peccoz P, Ferrari C et al. Plasma thyrotropin resoponse to thyrotropin-releasing hormone in patients with pituitary and hypothalamic disorders. J Clin Endo and Metab 1973;37:595–601. 488 Faglia G, Bitensky L, Pinchera A et al. Thyrotropin secretion in patients with central hypothyroidism: evidence for reduced biological activity of immunoreactive thyrotropin. J Clin Endo and Metab 1979;48:989–998. 489 Illig R, Krawczynska M, Torresani T, Prader A. Elevated plasma TSH and hypothyroidism in children with hypothalamic hypothyroidism. J Clin Endo and Metab 1975;41:722–728. 490 Snyder PJ, Jacobs LS, Rabello MM et al. Diagnostic value of thyrotropinreleasing hormone in pituitary and hypothalamic disorders. Ann Int Med 1974;81:751–757. 491 Bartalena L, Martino E, Falcone M et al. Evaluation of the nocturnal serum thyrotropin (TSH) surge, as assessed by TSH ultrasensitive assay, in patients receiving long term L-thyroxine suppression therapy and in patients with various thyroid disorders. J Clin Endo and Metab 1987;65:1265–1271. 492 Caron PJ, Nieman LK, Rose SR, Nisula GC. Deficient nocturnal surge of thyrotropin in central hypothyroidism. J Clin Endo and Metab 1986;62: 960–964. 493 Hayashizaki Y, Hiraoka Y, Tatsumi K et al. DNA analyses of five families with familial inherited thyroid stimulating hormone (TSH) deficiency. J Clin Endo and Metab 1990;71:792–796. 494 Docker BM, Pfaffle RW, Pohlenz J, Andler W. Congenital central hypothyroidism due to a homozygous mutation in the thyrotropin b-subunit gene follows an autosomal recessive inheritance. J Clin Endo and Metab 1998;83:1762–1765. 495 Rogol AD, Kahn CR. Congenital hypothyroidism in a young man with growth hormone, thyrotropin, and prolactin deficiencies. J Clin Endo and Metab 1976;39:356–363. 496 Winter JSD, DeGroot GW, Fairman C. Idiopathic sexual precocity in a boy with growth hormone, prolactin, and thyrotropin deficiencies. J Clin Endo and Metab 1974;39:356–363. 497 Wit JM, Drayer NM, Jansen M et al. Total deficiency of GH and prolactin and partial deficiency of thyroid stimulating hormone in two Dutch families: a new variant of hereditary pituitary deficiency. Horm Res 1989;32:170–177. 498 Yoshimoto M, Kinoshita E, Baba T et al. A case of severe pituitary dwarfism associated with prolactin and thyroid stimulating hormone deficiencies. Acta Paediatr Scand 1990;79:1247–1251. 499 Behringer RR, Mathews LS, Palmiter RD. Dwarf mice produced by genetic ablation of growth hormone expressing cells. Genes and Dev 1988;2:453–461. 500 Wu W, Cogan JD, Pfaffle RW et al. Mutations in PROP-1 cause familial combined pituitary hormone deficiency. Nature Genet 1998;18:147–149. 501 Fofanova O, Takamura N, Kinoshita E et al. Compound heterozygous deletion of the PROP-1 gene in children with combined pituitary hormone deficiency. J Clin Endo and Metab 1998;83:2601–2604. 502 Fluck C, Deladoey J, Rutishauser K et al. Phenotypic variability in familial combined pituitary hormone deficiency caused by a PROP-1 gene mutation resulting in the substitution of Arg–>Cys at codon 120 (R120C). J Clin Endo and Metab 1998;83:3727–3734. 503 Foley Jr TP, Owings J, Hayford JT, Blizzard RM. Serum thyrotropin responses to synthetic thyrotropin-releasing hormone in normal children and hypopituitary patients. J Clin Invest 1972;51:431–437. 504 Karlberg B, Almqvist S. Clinical experience with thyrotropin-releasing hormone (TRH) stimulation test in patients with thyroid, pituitary and hypothalamic disorders. Acta Endocrinol (Copenh.) 1973;72:697–713. 505 Schalch DS, Gonzales-Barcena D, Kastin AJ et al. Abnormalities in the release of TSH in response to TRH in patients with disorders of the pituitary, hypothalamus and basal ganglia. J Clin Endo and Metab 1972;35:609–615. 506 Costom BH, Grubach MM, Kaplan SL. Effect of thyrotropin-releasing factor on serum thyroid stimulating hormone: an approach to distinguishing

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hypothalamic from pituitary forms of idiopathic hypopituitary dwarfism. J Clin Invest 1971;50:2219–2225. Hall R, Ormston BJ, Besser GM, Cryer RJ. The thyrotropin-releasing hormone test in disease of the pituitary and hypothalamus. Lancet 1972;1:759–763. Mclaren EH, Hendricks S, Pimstone BL. Thyrotropin response to intravenous thyrotropin-releasing hormone in patients with hypothalamic and pituitary diseases. Clin Endocrinol (Oxf.) 1974;3:113–122. Mitsuma T, Shenkman L, Suphavai A, Hollander CS. Hypothalamic hypothyroidism: diminished thyroidal response to thyrotropin-releasing hormone. Am J Med Sci 1973;265:315–319. Suter SN, Kaplan SL, Aubert ML, Grumbach MM. Plasma prolactin and thyrotropin and the response to thyrotropin-releasing factor in children with primary and hypothalamic hypothyroidism. J Clin Endo and Metab 1978;47:1015–1020. Smallridge RC. Thyrotropin-secreting pituitary tumors. In: Mazzaferri EL, Samaan NA, eds. Endocrine Tumors. Boston: Blackwell Scientific Publications, 1993. Samuels MH, Wood WM, Gordon DF et al. Clinical and molecular studies of a thyrotropin-secreting pituitary adenoma. J Clin Endo and Metab 1989;68:1211–1215. Gershengorn MC, Weintraub BD. Thyrotropin-induced hyperthyroidism caused by selective pituitary resistance to thyroid hormone: a new syndrome of “inappropriate secretion of TSH”. J Clin Invest 1975;56:633–642. Weintraub BD, Gershengorn MC, Kourides IA, Fein H. Inappropriate secretion of thyroid-stimulating hormone. Ann Int Med 1981;95:339–351. Faglia G, Beck-Peccoz P, Piscitelli G, Medri G. Inappropriate secretion of thyrotropin by the pituitary. Horm Res 1987;26:79–99. Gittoes NJ, McCabe CJ, Verhaeg J et al. An abnormality of thyroid hormone receptor expression may explain abnormal thyrotropin production in thyrotropin-secreting pituitary tumors. Thyroid 1998;8:9–14. McDermott MT, Ridgway EC. Thyroid hormone resistance syndromes. Am J Med 1993;94:424–432. Refetoff S, Weiss RE, Usala SJ. The syndromes of resistance to thyroid hormone. Endocrine Rev 1993;14(3):348–399. McDermott MT, Ridgway EC. Central hypothyroidism. Endocrinol Metab Clin NA 1998;27:187–203. Refetoff S, DeWind LT, DeGroot LJ. Familial syndrome combining deafmutism, stippled epiphyses, goiter, and abnormally high PBI: possible target organ refractoriness to thyroid hormone. J Clin Endo and Metab 1967;27:279–294. Refetoff S. Syndromes of thyroid hormone resistance. American Journal of Physiology 1982;243:E88–E98. Pohlenz J, Weiss RE, Macchia PE et al. Five new families with resistance to thyroid hormone not caused by mutations in the thyroid hormone receptor b gene. J Clin Endo and Metab 1999;84:3919–3928. Refetoff S, Salazar A, Smith TJ, Scherberg NH. The consequences of inappropriate treatment because of failure to recognize the syndrome of pituitary and peripheral tissue resistance to thyroid hormone. Metabolism 1983;32:822–834. Lazar MA, Chin WW. Nuclear thyroid hormone receptors. J Clin Invest 1990;86:1777-1782. Usala SJ, Bale AE, Gesundheit N et al. Tight linkage between the syndrome of generalized thyroid hormone resistance and the human c-erbAb gene. Mol Endocrinol 1988;2:1217–1220. Chatterjee VKK, Nagaya T, Madison LD et al. Thyroid hormone resistance syndrome: inhibition of normal receptor function by mutant thyroid hormone receptors. J Clin Invest 1991;87:1977–1984. Parrilla R, Mixson AJ, McPherson JA et al. Characterization of seven novel mutations of the c-erbAb gene in unrelated kindreds with generalized thyroid hormone resistance: evidence for two “hot spot” regions in the ligand binding domain. J Clin Invest 1991;88:2123–2130. Yoh S, Chatterjee VKK, Privalski ML. Thyroid hormone resistance syndrome manifests as an aberrant interaction between mutant T3 receptor and transcriptional corepressor. Mol Endocrinol 1997;11:470–480. Liu Y, Takeshita A, Misiti S et al. Lack of coactivator interaction can be a mechanism for dominant negative activity by mutant thyroid hormone receptors. Endocrinology 1998;139:4197–4202. Weiss RE, Hayashi Y, Nagaya T et al. Dominant inheritance of resistance to thyroid hormone not linked to defects in the thyroid hormone receptors a or b genes may be due to a defective cofactor. J Clin Endo and Metab 1996;81:4196–4203. Sakurai A, Miyamoto T, Refetoff S, DeGroot LJ. Dominant negative transcriptional regulation by a mutant thyroid hormone receptor b in a family

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