PNH and the GPI-Linked Proteins

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Paroxysmal Nocturnal Hemoglobinuria and the Glycosylphosphatidylinositol-Linked Proteins

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Paroxysmal Nocturnal Hemoglobinuria and the GlycosylphosphatidylinositolLinked Proteins Edited by NEAL S. YOUNG JOEL MOSS National Heart, Lung and Blood Institute National Institutes of Health Bethesda, Maryland

San Diego San Francisco New York Boston London Sydney Tokyo

Cover photo credit.: Photo courtesy of Satyajit Mayor. For more information, please see Figure 10-1A on page 225. This book is printed on acid-free paper. 䊊 앝 Copyright © 2000 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Requests for permission to make copies of any part of the work should be mailed to: Permissions Department, Harcourt Inc., 6277 Sea Harbor Drive, Orlando, Florida 32887-6777

Academic Press A Harcourt Science and Technology Company 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www.apnet.com

Academic Press 24-28 Oval Road, London NW1 7DX, UK http://www.hbuk.co.uk/ap/ Library of Congress Catalog Card Number: 99-65876 International Standard Book Number: 0-12-772940-2 PRINTED IN THE UNITED STATES OF AMERICA 00 01 02 03 04 05 QW 9 8 7 6 5 4 3 2 1

Contents

Contributors Preface

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1 A Brief History of PNH Wendell Rosse

The Diagnosis of PNH Summary 15 References 16

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2 Genetics of PNH Lucio Luzzatto and Khe´doudja Nafa

Biochemical Genetics of PNH in Somatic Cells The PIG-A Gene 23

21

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Contents

The PIG-A Protein 24 Detecting Mutations in the PIG-A Gene 27 Germ Line PIG-A Mutations 32 The Spectrum of Somatic Mutations in the PIG-A Gene 33 PIG-A Mutations without PNH 40 Structure–Function Relationships 41 Correlation between Phenotypes and Genotypes 42 Comparison of the Spectrum of Mutations in Three X-Linked Genes: PIG-A, FVIII, and G6PD 42 Conclusion 43 References 44

3 Hemolysis in PNH Charles J. Parker

Introduction 49 The Classical Pathway of Complement 55 The Alternative Pathway of Complement 61 Phenotypic Mosaicism 64 Functional Basis of the Abnormal Sensitivity of the Erythrocytes of PNH to Complement-Mediated Lysis 68 Treatment of Hemolysis 83 Summary and Conclusions 86 References 89

4 Thrombotic Complications in PNH Elaine M. Sloand and Neal S. Young

Introduction 101 Incidence of Thrombosis in PNH 102 Factors Associated with Venous Thrombosis 103 Platelet Abnormalities in PNH 104 Role of Red Cell and Platelet Microvesicles 107 Urokinase Activator Receptor (uPAR) and Cell-Mediated Fibrinolysis 108

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Contents

Treatment of Thrombosis in PNH Summary 109 References 110

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5 Bone Marrow Failure in PNH Daniel E. Dunn, Johnson M. Liu, and Neal S. Young

Introduction 113 Pathophysiology of Marrow Failure in PNH 115 An Immune Model of PNH Pathogenesis 120 Clinical Course and Treatment of Marrow Failure in 127 PNH Summary 130 References 131

6 Animal Models of PNH Taroh Kinoshita, Monica Bessler, and Junji Takeda

Introduction 139 Mouse Piga Gene 141 In Vitro Differentiation of PIGA(⫺) ES Cells 143 Chimeric Mice Bearing PIGA(⫺) Hematopoietic Stem 149 Cells Hematopoiesis by Cells from Patients with PNH in SCID Mice 154 Conclusions and Perspectives 155 References 155

7 The Function of GPI-Anchored Proteins Ian Okazaki and Joel Moss

Introduction 159 GPI-Linked Proteins and Cell Signaling

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Contents

Role of GPI-Anchored Proteins in Insulin Action GPI-Anchored Proteins Function as Receptors GPI Anchors and Membrane Structure 168 Summary 170 References 171

166 166

8 GPI in Lower Animals Louis Schofield

Introduction 179 Structures of Protozoal GPIs 180 Evidence for GPI-Mediated Signaling in Mammalian Cell Biology 182 Signal Transduction in Host Cells by Protozoal GPIs 185 Parasite GPIs as Functional Toxins and Pathogenicity Factors 187 Regulation of Macrophage Function by Glycoconjugates of Leishmania 188 Regulation of Host Cells by Ceramide-Containing GPIs of T. cruzi 189 Insulin-Mimetic Activities of Protozoal GPIs 189 CD1d-Restricted NK T Cells and Immunity to GPI-Anchored Proteins 190 Conclusions 191 References 192

9 Synthesis of the GPI Anchor Daniel Sevlever, Rui Chen, and M. Edward Medof

Introduction 199 GPI Biosynthetic Pathway 200 Mammalian and Yeast GPI-Anchor Defective Mutants 203 GPI-Anchor Addition to Proteins 207 Biological Implications of GPI Anchoring Conclusions 213 References 214

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10 Functional and Structural Organization of GPI-Anchored Proteins in Cellular Membranes Satyajit Mayor

Introduction 221 GPI-Anchor Function and the ‘‘Raft’’ Hypothesis 221 Lateral Heterogeneity in Cell Membranes 222 Detergent-Resistant Membranes and Their Connection with 223 Rafts Evidence for Rafts in Living Cell Membranes 227 Involvement of Rafts in Biosynthetic and Endocytic Sorting 230 Rafts and Signaling by GPIAPs 232 In Conclusion 233 References 233

11 Structure and Function of GPI-Specific Phospholipases Martin G. Low

Introduction: GPI Anchors and Phospholipases 239 Biochemical Properties of Plasma GPI-PLD 245 Distribution of GPI-PLD in Tissues and Cells 252 Physiological Role of GPI-PLD 257 References 262

Appendix: Sequence of the Coding Region of the Human PIG-A Gene Index

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Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Monica Bessler (139) Division of Hematology, Department of Internal Medicine, Washington University School of Medicine, St. Louis, Missouri 63110. Rui Chen (199) Institute of Pathology, Case Western Reserve University, Cleveland, Ohio 44106. Daniel E. Dunn (113) Hematology Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892. Taroh Kinoshita (139) Department of Immunoregulation, Research Institute for Microbial Diseases, Osaka University, Osaka 565-0871, Japan. Johnson M. Liu (113) Hematology Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892. Martin G. Low (239) Department of Physiology and Cellular Biophyiscs, Columbia University, New York, New York 10032. Lucio Luzzatto (21) Department of Human Genetics, Memorial Sloan-Kettering Cancer Center, New York, New York 10021. Satyajit Mayor (221) National Centre for Biological Sciences, Hebbal, Bangalore 560 065, India. M. Edward Medof (199) Institute of Pathology, Case Western Reserve University, Cleveland, Ohio 44106. Joel Moss (159) Pulmonary-Critical Care Branch, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892. Khe´doudja Nafa (21) Department of Human Genetics, Memorial SloanKettering Cancer Center, New York, New York 10021. Ian Okazaki (159) Pulmonary-Critical Care Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892. Charles J. Parker (49) The Veterans Affairs Medical Center, Salt Lake City, Utah 84148; and Department of Medicine, Division of Hematology, University of Utah School of Medicine, Salt Lake City, Utah 84103. xi

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Contributors

Wendell Rosse (1) Duke University Medical Center, Durham, North Carolina 27710. Louis Schofield (179) Walter and Eliza Hall Institute of Medical Research, Royal Melbourne Hospital, Parkville, Victoria 3050, Australia. Daniel Sevlever (199) Department of Research, Mayo Clinic, Jacksonville, Florida 32224. Elaine M. Sloand (101) National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892. Junji Takeda (139) Department of Environmental Medicine, Osaka University, Osaka 565-0871, Japan. Neal S. Young (101, 113) National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892.

Preface

Paroxysmal Nocturnal Hemoglobinuria and the GlycosylphosphatidylinositolLinked Proteins reflects the recent fortunate coincidence of breakthroughs in basic science laboratories that impact our understanding of disease and of insights derived from the study of human illness that inspire such experiments. This book critically reviews the clinical character and underlying biology of paroxysmal nocturnal hemoglobinuria (PNH). Early studies of trypanosomes, a parasitic scourge of humans, led to the discovery of the glycosylphosphatidylinositol (GPI) anchor and eventually to the association of the red cell defect in PNH patients with the mammalian family of GPI-linked proteins. The recognition of a deficiency in a distinctive class of cell-surface proteins and of a single acquired genetic defect in hematopoietic stem cells has allowed definition of the genetic and biochemical nature of this strange disease. After more than a century of observation and study, we now understand how severe red blood cell destruction, life-threatening thrombosis, and aplastic anemia can be combined in a single patient, and the specific lesion responsible for intravascular hemoloysis has been defined. But many provocative questions still remain, ranging from the physiologic roles of GPI-anchored proteins and membrane rafts in the cell to the relationship between a protein defect and profound marrow failure. The authors of this volume are major investigators in biochemistry, molecular biology, and clinical medicine. As editors, we are especially appreciative of their willingness to prepare original syntheses for this monograph, and we hope that the readers will be educated and stimulated by their contributions. Neal S. Young Joel Moss

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1 A Brief History of PNH Wendell Rosse Duke University Medical Center, Durham, North Carolina 27710

Paroxysmal nocturnal hemoglobinuria (PNH) was one of the first discrete hematological entities to be described, undoubtedly due to the dramatic content of its primary symptom—hemoglobinuria. The passage of red, dark brown, purple, or black urine is readily noted by the patient and induces a distinct impression on the physician. The Hippocratean writings contain numerous allusions to ‘‘black urine’’ and scholars have debated whether they are describing blood or hemoglobin in the urine or simply concentrated urine, because most of the patients were moribund and most likely poorly hydrated. One word used (애␧␭움␷ 움) means specifically black or very dark, while another (앟움␹␷␴, which has the general meaning ‘‘thick’’) is used to describe ‘‘concocted’’ urine. The latter is clearly concentrated urine of the dehydrated (a not infrequent condition in Hippocrates’ case reports), leaving some doubt of the meaning of the former. However, a nocturnal or morning pattern of dark urine is not described, leading to the conclusion that Hippocrates probably is describing hemoglobinuria from malaria (many of the patients were febrile) or from some other cause other than PNH. In the Middle Ages, up to about the 17th century, close observation of the urine was a primary diagnostic test of the patient’s condition; textbooks detail what should be observed and the presumed causes for the changes seen. (Treatises on distinguishing horse urine from human urine were published because apparently the expertise of the physician was tested by substituting it for the urine of the patient.) Although dark black and red urine are described, the nightly pattern of PNH is not. The first case report which might be of a patient with PNH was that by Charles Stewart, a Scottish surgeon practicing in Archangel, the northernmost PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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port in Russia, who reported a patient with ‘‘periodical discharge of blood from the urethra’’ [1]. These crises of dark urine lasted about 3 days and were accompanied by pain in the lower back and abdomen, torpor, and a sense of fullness in the head, symptoms commonly seen in the hemoglobinuric crisis of PNH. Although a nocturnal pattern is not clearly described, the dark urine appeared intermittently and then abruptly ceased. No mention is made of exposure to cold as an inciting cause; in Archangel, this might have been difficult to determine. This probably represents the first reported case of PNH even though the details are very sketchy. The first certain description of PNH is that of William Gull (1816–1890), a leading physician at Guy’s Hospital, London in the mid- and late-19th century. He wrote a case report notable for its length and completeness and for its lack of any references [2]. He called the entity ‘‘intermittent haematinuria’’ because he recognized that the urine was ‘‘bloody-looking’’ but did not contain cells, clearly distinguishing it from hematuria. The patient was in the hospital from February 2 to March 26 (probably of 1865) and an accurate description of the state of his urine was reported for each day, using an arbitrary color scale ranging from indigo through mahogany to amber. Gull noted that the ‘‘haematinuria’’ occurred in episodes of 2–3 days and that the urine of the early morning was more affected than that later in the day. The patient had been employed as a leather-dye worker and had been exposed to cold and wet and Gull assumed that this may have been an ‘‘exciting’’ cause for the syndrome. In making this conclusion, he may have been influenced by reports of hemoglobinuria that had been published the previous year. Dr. George Harley, an assistant physician at University College Hospital, London, described two patients with ‘‘haematuria’’ who also had sudden and intermittent bouts of dark urine [3]. He distinguished this from ordinary hematuria as the latter had blood cells in the urine whereas his patient’s urine did not. One patient clearly had paroxysmal cold hemoglobinuria and the other may have had malaria, having recently been posted in the army to Walcheren in the sodden coastal regions of Holland. Several more cases of cold-induced hemoglobinuria were reported at the same meeting [4], suggesting that this condition was not rare. Six years later, paroxysmal cold hemoglobinuria (PCH), was linked to syphilis. Over the next few years, many more cases of hemoglobinuria were reported; most were probably PCH, but some may have been PNH, including three reported around 1880. In St. Petersburg Stolnikow described a patient with episodes of hemoglobinuria, sometimes quite severe, which did not seem to be related to exposure to cold. Because he collected the urine in 24-h aliquots, he failed to note a nocturnal pattern. He did have the insight to blame the pigmenturia on the destruction of the red cells, rather than on some abnormality of the kidney, as was most commonly done up to that time. The pattern of nocturnal hemoglobinuria was more accurately described by Lepine, a physician in Lyon, France [5]: As quoted by Crosby [6], he wrote: ‘‘It is only at night that the urine was blood-colored . . . Around 11:00 o’clock or midnight, the urine was blood-colored and not the other specimens.’’ He thought that the kidneys were at fault and that this represented a special case of hematuria in which the red cells were lysed during passage through the kidney or urinary

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1. A Brief History of PNH

William Withey Gull William Withey Gull was born in 1816 in Colchester and grew up in Thorpe-le-Soken, a small villiage that was owned by Thomas Guy’s Foundation. Through this association, he came to the attention of Benjamin Harrison, treasurer at Guy’s Hospital, who brought him to London at age 18. There he attended school for 3 years and worked at Guy’s Hospital before entering medical school. Four years later, he graduated with an MB from the University of London and 5 years after that (in 1846) he was awarded the MD with a gold medal in medicine. After holding various posts at Guy’s, he was given quarters in the hospital compound to set up practice; in this, he was successful from the beginning because, as one of his fellow physicians said, ‘‘(He) always gave satisfaction.’’ In 1846, he was appointed the first chair in physiology at Guy’s and by 1858, he had

been appointed full physician. His fame was greatly increased when he was called in (probably by mistake) to help in the care of the Prince of Wales in 1871 who was suffering from typhoid fever. The Prince recovered and Gull was made baronet. He wrote widely in medicine, including one of the first descriptions of anorexia nervosa, several on paraplegia, brain abscess, Bright’s disease (glomerulonephritis), rheumatic fever, ‘‘fatty stools from disease of the mesenteric glands,’’ and hypothyroidism. His classic article ‘‘On a Cretinoid State Supervening in Adult Life in Women’’ described myxedema so well that his name is often attached eponymously to this disorder. His article on ‘‘intermittent haematinuria’’ was his only writing on hematological subjects. He died in 1890, an honored and very wealthy physician.

The first clear account of a case of PNH was reported in 1866 by William (later Sir William) Gull [2]. Although the clinical findings were accurate, Gull had little insight into the pathophysiology of the disorder.

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tract. He did recognize that there were probably two kinds of hemoglobinuria, as his patient’s pigmenturia was not brought on by exposure to cold. The classic description of PNH is clearly that of Paul Stru¨bing, a young assistant physician in the clinic of Professor Mosler in Griefswald in eastern Germany [7]. He described a 29-year-old craftsman who, while in the Army, had noted dark red to black urine episodically, especially in the morning. The patient claimed that the disorder was service connected, as he had been subjected to adverse environmental conditions while in the Army. While having an episode, he noted weakness, fatigue, and easy exhaustion as well as pain, particularly in the area of the spleen; at the onset of an attack, there was shortness of breath, palpitation of the heart, a glitter before his eyes, buzzing in the ears, and vertigo. The kidney area was tender, as was the spleen; less tenderness was noted in the liver. Stru¨bing noted most of the symptoms that PNH patients have during an attack. Many patients have a feeling of excessive fatigue, clearly related to the hemolytic episode. Some have esophageal spasm, giving them shortness of breath. A symptom that Strubing did not describe was the impotence that almost all men have during an attack; such things were probably not easily discussed in 19th century Germany. Stru¨bing was most interested in factors that might precipitate an attack. Exposure to cold appeared to play no part, as the attacks were as common in summer as in winter. The patient was able to go into severe cold in light clothing and not have an attack. When ether was allowed to evaporate from his body,

Paul Stru ¨bing Paul Stru¨bing was born in Pyritz, Germany in 1852 and received his degree as doctor of medicine in 1876. He originally worked in the department of ophthalmology but turned to internal medicine and was made lecturer in 1882,

2 years after the appearance of his epochal papers. He became full professor of internal medicine in 1900. He wrote on many subjects, particularly related to otorhinolaryngology, but never again on any hematologic theme.

The account by Stru¨bing in 1882 [7] gave an even clearer clinical description and advanced some hypotheses about the cause of the symptoms. By this time, hemoglobinuria was well known and he propounded a careful differential diagnosis to show that his patient did not have march hemoglobinuria or paroxysmal cold hemoglobinuria.


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thus cooling the skin, no attack ensued. The patient did not have march hemoglobinuria, which had just been described by Fleischer [8] in a young Prussian army recruit who passed dark urine immediately following a ‘‘strenuous field exercise,’’ presumably in the wrong kind of footwear [9]. On the other hand, Stru¨bing’s patient did note dark urine after heavy work or exercise, not immediately but rather later that same night. Too much beer and ‘‘disagreeable emotions’’ were also cited as causes of episodes. The patient surreptitiously took iron salts and had a very severe attack, the explanation for which was only uncovered 89 years later [10]. (The repletion of iron causes a burst of erythropoiesis which produces a large number of cells susceptible to hemolysis.) Curiously, Stru¨bing did not mention infections (which Gull had noted as a cause of attacks) as an etiology. Stru¨bing was particularly intrigued by the nocturnal component of the hemoglobinuria. He recognized that the source of the pigment was the red cells (because the plasma also turned red during an attack, which would not occur if the destruction were restricted to the kidneys), and he thought that the red cells were abnormal in some way, causing them to ‘‘disintegrate.’’ The professor of physiology at Greifswald at the time was Leonhard Landois, a relatively young, but widely recognized scientist who had investigated the transfusion of blood in animals and who had apparently a great influence on young Dr. Stru¨bing (17 of 54 references given in his paper are the work of Professor Landois). It is not clear which of the two originally theorized that the hemolysis of the red cells might be due to the accumulation of CO2 during sleep, thus acidifying the blood. They knew that normal red cells could be lysed in vitro by acid and thought perhaps the cells of the patient were somehow more susceptible to acid lysis. Stru¨bing gave the patient acid in an attempt to induce an attack but was not successful; Crosby [6] claims he gave too little, but it is doubtful that enough could be given and have the patient survive. Nevertheless, this seminal thought resurfaced several times and ultimately played a crucial role in the understanding of the disease. Stru¨bing noticed hemosiderin in the urine of his patient, although he did not identify it chemically. Landois had also noted this same pigment in the urine of animals who had reacted to xenogeneic transfusions, and he recognized it as a product of the red cells that had been destroyed in the immune reaction following transfusion. He suggested to Stru¨bing that the red cells in PNH might be hemolyzed as a result of being attacked as ‘‘foreign’’ cells; this was in the predawn of immunology and adumbrates the eventual finding that the hemolysis in both situations is due to immunological mechanisms. (These experiments in xenotransfusion in animals preceded the similar but more famous ones of Ehrlich [11] by about 20 years.) In his experiments on transfusion in animals, Landois had noted extensive microscopic thrombosis in the animals that had received xenogeneic blood and he probably suggested to Stru¨bing that some of the symptoms that his patients described could be due to a similar cause. Stru¨bing’s patient did have an enlarged spleen, usually a sign of thrombosis in the portal drainage area, but there was no other evidence of thrombosis. The role of thrombosis in the pathogenesis of the clinical description of PNH came much later.

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Stru¨bing’s paper is a masterful work that shows both clear clinical description and solid (if not always correct) scientific thought. But it was essentially ignored in the numerous clinical descriptions of the next 70 years, until resurrected by Crosby in 1951 [6]. The next major description of PNH came some 30 years after Stru¨bing’s. Hijmans van den Bergh was a physician at the Coolsingel Hospital in Rotterdam who, in 1911, described a 47-year-old man who had had chronic hemoglobinuria accompanied by anemia of variable degree [12]. By this time, the concept of hemolytic anemia had been well established by Chauffard and Vincent, and Chauffard published a case report of PNH in 1908, calling it ‘‘cas d’icte`re hemolytique’’ [13] (a case of hemolytic jaundice); this same patient was later described by Marchiafava. Van den Bergh recognized that his patient also had this type of disorder. By this time, considerably more knowledge and methodology were available than in Stru¨bing’s time. The science of immunology was in its late childhood, following on Metschnikoff and Ehrlich. Bordet [14] had outlined the elements of the humoral immune system with his famous paragraph describing the ‘‘principles’’ in serum that are necessary for hemolysis—one (now called antibody) was not thermolabile and was found in the blood only after sensitization of the animal by antigen. The other, which was thermolabile, was present in normal serum and augmented the activities of the first—for this latter action, Bordet called this principle ‘‘complement.’’ Ehrlich had clearly demonstrated how these two ‘‘principles’’ interacted in bringing about hemolysis [11]. Finally, Donath and Landsteiner had shown the hemolytic action of antibody and complement in paroxysmal cold hemoglobinuria by in vitro lysis of red cells by the patient’s serum in a bithermic reaction [15, 16], although it is not entirely clear that they understood the mechanism for this striking reaction [17]. Van den Bergh scrupulously used the best biological methods to investigate the nature of his patient’s illness. The cells of the patient did not hemolyze abnormally in hypotonic saline solutions, the test of Donath and Landsteiner for PCH was negative, and there was no ‘‘hemolysine’’ in the serum by the usual tests. On the other hand, when tests for hemolysis were done under conditions of acidification (by placing the reaction mixture under an atmosphere of CO2), the patient’s cells were hemolyzed both by his own serum and more so by the serum of a blood group compatible normal donor. The lytic principle was clearly different from antibody, as it was present in normal serum, and it was destroyed by heating to 50⬚C for 2 h, like complement. However, lysis was not restored by adding a ‘‘petite quantite´’’ of neat human or guinea pig serum. (Too little was added and thus the role of complement in the lysis of the red cells in PNH was missed.) Over the next 20 years, several more clinical descriptions appeared, the most famous being those of Marchiafava and his pupil, Micheli. The first report by Marchiafava [18] was made on the same patient that Chauffard and Troisier had previously published, a fact known but not referenced by the authors. Neither Marchiafava or Micheli added much to the previous clinical descriptions, except for the fact that patients had ‘‘perpetual hemosiderinuria,’’ even when they did not have hemoglobinuria. Nevertheless, the reports were well publicized and the other prior cases were forgotten; their name was (and still is) attached to the

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Hijmans van den Bergh A. A. H(ijmans) van den Bergh was born in Rotterdam in 1869. He graduated from Leiden in 1896 and, in 1900, he went to Breslau in Germany, the center of the chemical industry. He worked with the pediatrician, Czerny, but was undoubtedly influenced by Emil Fischer and his work on porphyrins. He returned to Rotterdam to practice where he encountered the patient with PNH described in his paper. In 1912, he succeeded Wenckebach as Professor of Medicine at Groningen when the latter went to Strasburg. In 1918, he was appointed Professor of Medicine at Utrecht where he con-

tinued his interest in prophyria and hemoglobinuria; he retired in 1938 at about the time of the publications on PNH of his pupil, Jordan. At his arrival in Utrecht in 1918, he became personal physician of the exiled German emperor, Wilhelm II, who lived in nearby Doom, and remained so until the death of the latter in 1941, even during the German occupation. Although he was Jewish, he was not interned by the Germans but was kept under house arrest until his death in 1943. He is still widely regarded as one of the pillars of Dutch medicine.

The first demonstration of the abnormality of the red cells in PNH was by Hijmans van den Bergh in 1911 [12]. He showed that the cells of the patient were hemolyzed by normal serum as well as by the patient’s own serum when it was acidified by exposure to carbon dioxide. Although he suspected complement was involved, he was not able to do the experiments to prove it. The potential of this reaction for diagnosis was ignored for 27 years.

condition. The first American (and the first English language report) was that of Giffen at the Mayo Clinic in 1922 who described the feeling of weakness, depression, and headache that accompanies the hemolytic episode in many patients [19]. It was not until 1928 that the name, paroxysmal nocturnal hemoglobinuria, as translated from the Latin, was first used by Enneking, a Dutch physician from Amsterdam, in a succinct report which differentiated the syndrome from other forms of hemoglobinuria and myoglobinuria [20]. Several other reviews were published in the 1930s. Hamburger and Bernstein [21], in a somewhat leisurely account, summarized the clinical data of 22 cases of PNH in the literature (they missed both Strubing and Gull’s accounts). They

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discussed hemoglobinuria in general, distinguishing the patients with PNH from those with myoglobinuria, paroxysmal cold hemoglobinuria, and march hemoglobinuria, as well as from patients with spherocytic hemolytic anemia. Chronicity, the occurrence of venous thromboses, and the inutility of splenectomy were emphasized; many attempts at modification of the hemoglobinuria were discussed but none seemed sovereign. The anatomic pathology was best described in a review by Bodley-Scott and his associates [22] in a review of 14 cases, including two of their own. They again stressed the occurrence of thromboses, including hepatic thrombosis, and changes that occurred in the kidneys due to the accumulation of large amounts of hemosideroin. Up to this time, only those patients with a pattern of nocturnal hemoglobinuria were recognized with PNH, as the diagnosis was made on purely clinical grounds. With specific diagnostic laboratory tests, it became clear that many patients had the disease but did not have hemoglobinuria. These diagnostic tests evolved from an attempt to understand the cause of the hemoglobinuria.

The Diagnosis of PNH By the late 1930s, the clinical syndrome of classic hemolytic PNH had been described, largely based on the occurrence of intermittent hemoglobinuria and ‘‘perpetual hemosiderinuria,’’ but little had been done to examine its causes. Then, almost simultaneously, three laboratories undertook experiments to explain why hemolysis occurred—Thomas Hale Ham in Boston, John Dacie in London, and F. L. J. Jordan in the Netherlands. The first to reach publication (in early 1938 [23]) was Jordan, who worked in the Clinique Universitaire of Dr. Hijmans van den Bergh in Utrecht. He had prepared his doctoral thesis on hemoglobinuria [24]. He compared the in vitro hemolytic reactions of PCH with those of ‘‘atypical hemoglobinuria, similar to the nocturnal hemoglobinuria of Enneking.’’ Complement was important, as in both cases, hemolysis was increased by lower pH. He clearly recognized, as had his mentor, that the red cells of the ‘‘atypical’’ patient were defective. The serum appeared to be normal, as no antibodies or other abnormal elements could be demonstrated but, because antibody (or ‘‘amboceptor’’) was the only way known for complement to be activated, he presumed that the cells had been sensitized in vivo. Ham’s initial report appeared in December 1938 [25]; it was short and did not reference any previous experimental work. A longer and more complete clinical paper appeared later the next year [26]. He argued that the nocturnal pattern of hemolysis could be due to acidification of the blood during sleep. When he gave sodium bicarbonate, there was a decrease in hemolysis, followed by a rebound; conversely ammonium chloride increased hemolysis. Placing the patient in a Drinker respirator so that the pH would not fall during sleep decreased the urine and plasma hemoglobin levels. These studies were reflected in vitro by a marked increase in hemolysis of the patient’s cells in his own or normal serum when the pH was reduced. Later that same year, Ham wrote his classic paper on the mechanism of lysis of the red cells in PNH [27]. He carefully examined the elements of immune

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Thomas Hale Ham Thomas Hale Ham was born in 1905 in Oklahoma but grew up in Yonkers, New York. He went to Dartmouth College, then to Cornell Medical School, followed by New York Hospital for his residency. He later went to Boston, to the Thorndike Memorial Laboratory, where he worked very successfully with Dr. William Castle, with whom he published his first paper on intrinsic factor in 1936. He struck out on his own investigations of PNH which he pursued

for the next few years. After serving in the Army during World War II, he returned to the Thorndike briefly but left in 1950 to lead the educational revolution at Western Reserve Medical School. He spent the rest of his life up to his retirement there and became widely respected as an educational innovator and a superb teacher. He retired to Vermont near his alma mater, and died in 1987 at the age of 81.

The report of Thomas Hale Ham in 1938 was one of three that appeared about the same time, demonstrating that the erythrocytes of patients with PNH were lysed by acidified serum. This provided a specific diagnostic test that identified patients who did not have the classical symptom of nocturnal hemoglobinuria.

hemolysis as they were understood and concluded that he could not demonstrate any evidence of antibody or of an unusual antigen on the PNH cells (but he could not rule out that the hemolytic reactions might be initiated by an antibody that could not be detected). He convincingly established that lysis was effected by complement by showing a dose–response relationship (limited because hemolysis disappears with even very little dilution of serum), and that inhibition or destruction of complement or components of complement abrogated the lytic reaction. He extensively examined the reaction and found that the difference in lysis between normal and PNH red cells depended upon the source of complement; most animal sources showed no difference but lysis was usually correlated to the use of human complement. We now know that the cellular inhibitors of complement are species-specific and not effective in inhibiting complement from heterologous sources. Ham found that PNH cells were more readily lysed when complement was activated by isologous antibodies, but he could not demonstrate any increase in complement fixation by PNH cells using classic tests. Dacie’s paper appeared in February 1939, 2 months after Ham’s [28] and also described hemolysis in acidified serum of red cells of patients with PNH.

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John Vivian Dacie John Vivian Dacie was born in 1912, the son of a clergyman, in Putney in southwestern London. He was educated at Kings College School in Wimbledon and Kings College and Kings College Hospital, receiving his medical degree in 1935; while there, he came under the influence of Janet Vaughn who sparked his lifelong interest in hematology. Shortly thereafter, he went to the Postgraduate Medical School (later the Royal Postgraduate Medical School) at Hammersmith Hospital as Pathologist. After a prolonged stint in the Royal Army, he returned to the Postgraduate Medical School and had a distinguished career until his retirement in

1975. He was (and is) keenly interested in the hemolytic anemias and his classic book Haemolytic Anaemias has gone to three editions, each a classic in its time; the final volume appeared in 1999 [96]. Although, as a pathologist, he did not directly care for patients, he was responsible for a number of seminal clinical papers about PNH and other hemolytic syndromes. For his contributions to medicine, he was knighted. He is almost as well known in the field of lepidoptery as in medicine. Since his retirement in 1977, he has continued to write and lives with his wife in Wimbledon.

Professor Sir John Dacie (shown here in his 87th year) authored his first contribution concerning PNH in 1938 (a description of a case with the demonstration of lysis of the erythrocytes by acidified serum) and his last 60 years later. He, more than any other single individual, has greatly contributed to the clinical and laboratory description of PNH.


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He noted the same sorts of reactions as the other two investigators but concluded that hemolysis was initiated by an elusive ‘‘lysin.’’ Ten years later, he published detailed results concerning initiation of the reaction, suggesting as a substance present in both normal and patients’ serum that was distinct from complement [29]; several years passed before this factor was understood to be part of a system of complement activation that does not depend upon antibody. Who should be credited for finding a test to diagnose PNH? Clearly, Jordan’s work has precedence. Stories abound about a rush to publication by Ham and by Dacie, who appear to have been working in parallel. On the other hand, the true discoverer was Hijmans van den Bergh, even though his work was ignored by all except his pupils; had he been more thorough and had he been recognized, we might have had another van den Bergh reaction in addition to that used in the analysis of serum bilirubin. In the end, a characteristic and diagnostic abnormality of the red cells of patients with PNH was identified—the unusual susceptibility of the red cells to lysis by complement. The acidified serum lysis test had the advantage of specificity; no other red cells were similarly lysed except in those of patients with the extremely rare congenital condition HEMPAS (hereditary erythroblastic multinuclearity with a positive acidified serum lysis test or congenital erythroblastic anemia, type II) [30]. In contrast to PNH, cells in this disease are not lysed by the patient’s own serum. The Ham or acidified serum lysis test did not detect very small populations of abnormal cells. Modifications to increase sensitivity, including adding thrombin to the mixture [31] and optimizing the concentration of Mg2⫹ [32], were modestly successful. Lysis by complement and dilutions of cold agglutinin antibodies [33] and by reduction in the ionic strength at which complement was reacted [34] increased the sensitivity, but lessened the specificity. Studies of Rosse and Dacie optimized conditions for detecting the complement-sensitive cells by maximally sensitizing cells with a complement-activating antibody and effecting lysis by limiting amounts of complement [35, 36]. With respect to lysis by complement, in most patients, two or three kinds of red cells were present: PNH I cells which appeared to be normal, PNH II cells which were somewhat sensitive to complement, and PNH III cells which were lysed by very low concentrations of complement. The reason for these differences became evident only 25 years later, but these results did suggest that PNH was a clonal disease, a conclusion that was substantiated by the finding that granulocytes and platelets also consist of complement-sensitive and insensitive cells [37, 38]. As knowledge of hematopoiesis grew over the next few years, it became apparent that PNH was the result of one or more defective hematopoietic stem cells that give rise to clones of abnormal blood cells [39]. The peculiar sensitivity of the red cells to lysis by complement played a small but important role in unraveling the complexities of the complement system. Complement had first been described as quiescent in the serum until activated by antibody, and no other mechanism of activation was recognized. In the early 1950s, Pillemer and his associates suggested that there was an alternative system for the activation of complement, involving a protein which they called ‘‘properdin’’ [40]. The identification of the components of complement had always depended upon inactivation or removal of components by various chemical and

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Wendell Rosse

physical means. The first component, which reacted with antibody, was thermolabile and required the presence of Ca2⫹. The second component depended upon Mg2⫹. The fourth component was inactivated by hydrazine and the ‘‘third’’ component did not require cations, was not thermolabile, and was not inactivated by hydrazine. Pillemer found that reaction of serum with zymosan, a complex polysaccharide derived from yeast cell walls, removed a component(s) which did not fit the definitions of the other known factors. Because antibodies to zymosan could not be detected in the serum, he proposed that complement had been activated by a ‘‘properdin pathway’’ that did not involve antibody and the first three components, C1, C4, and C2. The ‘‘establishment’’ in the world of complement did not believe Pillemer and claimed that all the reactions that he described could be explained by ‘‘naturally occurring’’ antibodies in the serum [41]. It became more and more difficult for him to find a journal that would accept his work and he became increasingly despondent in his frustration, eventually committing suicide. The critics of his work were, however, never able to explain the lysis of PNH red cells by acidified serum, as the reaction clearly did not involve antibody and ‘‘naturally occurring’’ heteroantibodies could not be evoked [42]. Eventually, when better immunochemical methods were developed, it became clear that Pillemer had been correct. The so-called ‘‘alternative pathway’’ of complement activation was unraveled and was shown to consist of components that were activated in the absence of antibody; lysis in acidified serum, and presumably also in the patient, was the result of the activation of this pathway [43]. This understanding came too late for Pillemer to receive his just desserts.

Thrombosis in PNH The occurrence of thrombosis in PNH had been noted in the early part of the 20th century but was not emphasized until the early 1950s. Crosby, in his monumental review, noted the clear relationship of thrombosis and PNH [44, 45]; nearly half (24/53) of the deaths he recorded were due to thrombosis. From this clue, he performed a series of experiments that suggested that activation of the coagulation system was responsible for initiation of hemolysis [46]. The addition of thrombin appeared to increase the lysis seen in the acidified serum lysis test [31] which he ascribed to destruction of a factor inhibitory of lysis in acidified serum; such a factor has not been identified. When the ‘‘properdin pathway’’ was found, increased lysis on the addition of thrombin was blamed on contamination of thrombin preparations with properdin [42], but Crosby was able to show that complete removal of properdin did not diminish the ability of thrombin to increase lysis of PNH cells in acidified serum [47]. Thrombin, which is a serine protease, might activate factor B or some other component of the alternative pathway, thus increasing the amount of lysis seen for a given activation of complement. However, the test never replaced acidified serum lysis (performed without the addition of thrombin) for the diagnosis of PNH. Over the next few years, specific clinical syndromes due to thrombosis of veins in unusual sites were described. The studies of Hartmann and his group emphasized the occurrence of thrombosis of the hepatic veins (Budd-Chiari


13

syndrome) in patients with PNH [48], now recognized as a fairly common and very serious complication of the disorder. Unusual sites of venous thrombosis (cerebral veins, dermal veins, and portal and splanchnic veins) were described. Epidemiological studies have shown about 40% of patients of European origin or ancestry have thromboses [49] and that their occurrence is a major adverse prognostic factor. Curiously, Asians with PNH have a much lower incidence of thrombosis (about 5–10%) [50, 51, 52], as do Mexicans, who are primarily of Asian origin through migration of the aboriginal Indian populations from that region [53]. PNH patients of south India, on the other hand, were found to have thrombotic complications [54]. The reason for the increased thrombosis in PNH is not clearly known. Intravascular release of membrane fragments arising from disruption of cells in the circulation might activate the procoagulant system, as could be shown in vitro, but proof was lacking. Once the platelets were shown to be abnormal in PNH, they were suspected of playing a role in the pathogenesis of the increased thrombosis. When platelets (and granulocytes) are attacked by complement, they protect themselves by removing the nascent C9 complexes by exovesiculation [55]; these vesicles are sites for the formation of prothrombinase complexes and are thus thrombogenic [56]. Because PNH platelets lack CD59, an important inhibitor of the formation of polymeric C9 complexes, the activation of even small amounts of complement on their surface results in a massive production of these microvesicles [57]. Further, the monocytes in PNH lack the urokinase–plasminogen activator receptor (uPAR) [58] which probably inhibits fibrinolysis when thrombosis has occurred. There may well be other factors that play a role in the interaction between two complex systems—complement and coagulation—that lead to thrombosis in PNH.

On the Trail of the Cellular Defect Stru¨bing suspected and Hijmans van den Bergh proved that the red cells were abnormal in PNH, but the biochemical nature of the abnormality remained elusive for many years. Clues came from studies of cellular enzymes, as these could be readily detected by their activity. Leukocyte alkaline phosphatase was found to be lacking in a population of neutrophils [59, 60] and acetylcholinesterase was deficient in the red cells of patients with PNH [61]. The abnormal erythrocyte population was the one lacking the enzyme, and complement sensitivity was inversely related to the concentration of the enzyme on the cell surface [62, 63]. However, no direct relationship could be established between the lack of acetylcholinesterase and the susceptibility of the cells to lysis by complement. Many attempts were made to reproduce the complement sensitivity of the PNH erythrocytes by incubation of normal cells with various agents including proteases [64] and sulfhydryl-reactive compounds [65–67]. These studies suggested that the abnormality probably involved proteins that required disulfide bonds for their activity, but did not directly lead to an elucidation of the basis for complement sensitivity. A major advance in the biochemistry of complement sensitivity came with the identification of decay-accelerating factor (DAF), a protein that regulates the action of complement on the cell surface [68–70]; credit is usually given to

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Anne Nicholson-Weller and her associates for this important discovery, although the studies of Hoffman several years earlier had indicated its presence. The protein has been well characterized and has been shown to act by disrupting or preventing the formation of the bimolecular amplification convertase complexes of both the classical (C4b2a) and alternative (C3bBb) pathways. Early studies using the methods of Hoffman failed to show a difference between normal and PNH red cells with respect to the activity of this protein [71], in part because guinea pig components of complement were used rather than human ones. However, the more careful studies of Nicholson-Weller [72] and of Pangburn and his associates [73] clearly demonstrated that abnormal cells in PNH lacked this activity, due to the absence of the protein rather than inhibition of its enzymatic activity. Lack of DAF was thought for a time to explain the sensitivity to complement of the PNH red cells because it could be shown that these cells fixed much more C3 for a given level of complement activation. On further investigation, however, some step beyond the fixation of C3 was also found to be abnormal, as much more lysis of PNH cells resulted from an equivalent fixation of that component [74]. This defect was due to the lack of another protein, variously called membrane inhibitor of reactive lysis (MIRL) [75], HRF20 [76], MACIF [77], protectin [78], and, most commonly, CD59. CD59 prevents the formation of polymeric complexes of C9, the complement component that ultimately penetrates the membrane and causes hemolysis. The absence of CD59 on the membranes of the red cells of PNH accounts for most of the sensitivity to complement [79]. The molecular basis of these several protein deficiencies was not obvious. They were clearly different proteins [80] and were derived from genes on different chromosomes. The solution came with the recognition that one of them, alkaline phosphatase, was affixed to the membrane by a complex glycosylphosphatidylinositol anchor [81, 82], rather than through a sequence of nonpolar amino acids. DAF [83] and CD59 [84] could also be demonstrated to be attached by the same mechanism, suggesting that a defect in the biosynthesis of the anchor might account for deficiency of all the proteins. This biosynthetic deficiency was soon described and the specific gene, PIG-A was cloned by Kinoshita and his associates; the abnormal hematopoietic cells of all PNH patients have mutations in the PIG-A gene [85, 86]. This remarkable series of observations and deductions began with clinical observations and led to biochemical and finally genetic definitions of the cause of the disease. Serendipity played an important role, but more important was the application in a logical manner of increasingly sophisticated techniques to an interesting problem.

The Relationship of PNH and Aplastic Anemia The absence of the glycosylphosphatidylinositol (GPI)-linked protein explains many of the clinical symptoms of PNH, but one manifestation, its relationship to aplastic anemia and hematopoietic deficiency, is not yet explained. Many early reports of aplasia in PNH suggested that the marrow became ‘‘aregenerative.’’ The clinical relationship between aplastic anemia and PNH became clear from the studies of Dacie and Lewis [87, 88]. They found that about 25% of patients who had PNH had significant bone marrow hypoplasia or had a history of it or


15

who had a history of significant bone marrow hypoplasia and that 15% of patients with aplastic anemia had a positive acidified serum lysis test; the association of PNH and aplastic anemia became even more evident when a relatively successful resolution of aplasia could be obtained by the use of antilymphocyte sera in its treatment [89, 90]. It was also determined experimentally and by observation that hematopoiesis was impaired even in patients who did not have a history of aplastic anemia. Many patients with apparently active hematopoiesis have thrombocytopenia to a variable degree [91]. Studies have shown that in the absence of splenomegaly, the survival of the platelets in the circulation is normal [92, 93]; the thrombocytopenia must be due to deficient production. This is also true for the granulocytes; neutropenia is not uncommon [91], but the survival of granulocytes in the circulation is normal [86]. Further, as Crosby [45] and Dacie and Lewis [87] noted, the level of reticulocytosis in PNH is less for the degree of anemia than is seen in other hemolytic syndromes. All these studies indicated that hematopoiesis was usually, if not always, impaired, despite the cellular appearance of the bone marrow biopsy. When cell culture methods became available, hematopoiesis could be directly assessed in vitro. Early studies showed that bone marrow from PNH patients, even those with apparently active hematopoiesis by marrow morphology, did not yield a normal number of colonies [94, 95]. Related to the problem of deficient hematopoiesis is the question of how the PNH clone, arising from a single cell, is able to encompass the major part of hematopoiesis. No convincing evidence, exists to indicate that the defective cells have a growth advantage of normal cells. Studies such as these led to the Young–Luzzatto hypothesis which, in summary, states: 1. Many normal individuals have a very small number of hematopoietic stem cells in which the PIG-A gene is mutated, but these cells are at a growth disadvantage and remain a tiny minority; 2. If the bone marrow is afflicted by an ‘‘aplastogenic’’ process (probably autoimmune), these cells are less affected than normal precursors and the PNH clone becomes evident and usually even dominant in the marrow. Thus, PNH would be the result of interaction between an acquired somatic defect in the hematopoietic precursors and an alteration in hematopoiesis by a (presumably) unrelated process. As in the past, the investigation of these factors will provide information not only about this disease but about human biology in general and other diseases in particular.

Summary The history of PNH shows many examples of the role of careful clinical observation and the application of the methods of science available at the time in unraveling this complex disorder. As with all science, it is characterized by a series of ‘‘platforms’’ which solidify understanding and which are built upon as more observations are made and better methods become available. Chauffard and Vincent wrote in 1909: ‘‘Nous ne sommes qu’au seuil de ces recherches expe´rimentales, et le progre`s actuel doit consister a` e´tudier scrupuleusement les faits cliniques par toutes les me´thodes biologiques dont nous disposons’’ (we are only at the threshhold of this experi-

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mental research and actual progress ought to consist in scrupulously studying the clinical facts by all the biological methods at our disposal)—an excellent guiding maxim. We are always at the threshhold of new experimental research, and our progress should consist of the scrupulous determination of the clinical facts, using all the biological and biochemical methods that we have at our disposal. Our knowledge is never complete.

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H., Cahn, J.-Y., and Gluckman, E. (1996). Paroxysmal nocturnal haemoglobinuria: long term follow-up and prognostic factors. Lancet 348, 573–577. Le, X. F., Yang, T. Y., Yang, X. Y., and Wang, X. M. (1990). Characteristics of paroxysmal nocturnal hemoglobinuria in China. Clinical analysis of 476 cases. Chin. Med. J. 103, 885–889. Fujioka, S., and Asai, T. (1989). Prognostic features of paroxysmal nocturnal hemoglobinuria in Japan. Acta Haematol. Jpn. 52, 1386–1394. Kruatrachue, M., Wasi, P., and Nanakorn, S. (1978). Paroxysmal nocturnal haemoglobinuria in Thailand with special reference to an association with aplastic anemia. Brit. J. Haematol. 39, 267–276. Gongora-Bianchi, R. A., Gonzalez-Martinez, P., Sosa-Munoz, J., Castro-Sansores, C., DelgadoLamas, J. L., Vasquez-Villegas, V., Rico-Balzadua, G., Herrera-Sanchez, P., Selva-Pallares, J., and Gonzalez-Llaven, J. (1997). Natural history of paroxysmal nocturnal hemoglobinuria in adolescents, adults, and children. The Mexican experience. Sangre 42, 171–177. Koduri, P. R., and Gowrishankar, S. (1992). Paroxysmal nocturnal haemoglobinuria in Indians. Acta. Haematol. 88, 126–128. Sims. P. J., Faioni, E. M., Wiedmer, T., and Shattil, S. J. (1988). Complement proteins C5b-9 cause release of membrane vesicles from the platelet surface that are enriched in the membrane receptor for coagulation factor Va and express prothrombinase activity. J. Biol. Chem. 263, 18205– 18212. Gilbert, G. E., Sims, P. J., Wiedmer, T., Furie, B., Furie, B. C., and Shattil, S. J. (1991). Plateletderived microparticles express high affinity receptors for factor VIII. J. Biol. Chem. 266, 17261– 17268. Wiedmer, T., Hall, S. E., Ortel, T. L., Kane, W. H., Rosse, W. F., and Sims, P. J. (1993). Complement-induced vesiculation and exposure of membrane prothrombinase sites in platelets of paroxysmal nocturnal hemoglobinuria. Blood 82, 1192–1196. Ploug, M., Plesner, T., Ronne, E., Ellis, V., Hoyer Hansen, G., Hansen, N. E., and Dano, K. (1992). The receptor for urokinase-type plasminogen activator is deficient on peripheral blood leukocytes in patients with paroxysmal nocturnal hemoglobinuria. Blood 79, 1447–1455. Beck, W. S., and Valentine, W. N. (1951). Biochemical studies on leucocytes. II. Phosphatase activity in chronic lymphatic leukemia, acute leukemia, and miscellaneous hematologic conditions. J. Lab. Clin. Med. 38, 245–253. Lewis, S. M., and Dacie, J. V. (1965). Neutrophil (leucocyte) alkaline phosphatase in paroxysmal nocturnal haemoglobinuria. Brit. J. Haematol. 11, 549–557. Auditore, J. V., Hartmann, R. C., Flexner, J. M., and Balchum, O. J. (1960). The erythrocyte acetylcholinesterase enzyme in paroxysmal nocturnal hemoglobinuria. Arch. Pathol. 69, 534–540. Kunstling, T. R., and Rosse, W. F. (1969). Erythrocyte acetylcholinesterase deficiency in paroxysmal nocturnal hemoglobinuria (PNH). A comparison of the complement-sensitive and insensitive populations. Blood 33, 607–616. Chow, F. L., Telen, M. J., and Rosse, W. F. (1985). The acetylcholinesterase defect in paroxysmal nocturnal hemoglobinuria: Evidence that the enzyme is absent from the cell membrane. Blood 66, 940–945. Yachnin, S., Laforet, M. T., and Gardner, F. H. (1961). pH dependent hemolytic systems. I. Their relationship to paroxysmal nocturnal hemoglobinuria. Blood 17, 83–96. Sirchia, G., Ferrone, S., and Mercuriali, F. (1965). The action of two sulfhydryl compounds on normal human red cells: Relationship to red cells of paroxysmal nocturnal hemoglobinuria. Blood 25, 502–508. Sirchia, G., and Dacie, J. V. (1967). Immune lysis of AET-treated normal red cells (PNH-like cells). Nature 215, 747–748. Stead, N. W., and Rosse, W. F. (1971). Quantitative studies on the sensitivity to complement lysis of red cells treated with thiol-reactive reagents. Blood 37, 454–462. Hoffman, E. M. (1969). Inhibition of complement by a substance isolated from human erythrocytes. I. Extraction from human erythrocyte stromata. Immunochemistry 6, 391–403. Hoffman, E. M. (1969). Inhibition of complement by a substance isolated from human erythrocytes. II. Studies on the site and mechanism of action. Immunochemistry 6, 405–419. Nicholson-Weller, A., Burge, J., Fearon, D. T., and Weller, P. F. (1982). Isolation of a human erythrocyte membrane glycoprotein with decay-accelerating activity for C3 convertases of the complement system. J. Immunol. 129, 184–189.


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71. Chua, C., Hoffmann, E. M., Adams, J. P., and Rosse, W. F. (1980). Inhibitors of complement derived from the erythrocyte membrane in paroxysmal nocturnal hemoglobinuria. Blood 55, 772–776. 72. Nicholson-Weller, A., March, J. P., Rosenfeld, S. I., and Austen, K. F. (1983). Affected erythrocytes of patients with paroxysmal nocturnal hemoglobinuria are deficient in the complement regulatory protein, decay accelerating factor. Proc. Natl. Acad. Sci. U.S.A 80, 5430–5434. 73. Pangburn, M. K., Schreiber, R. D., and Mu¨ller-Eberhard, H. J. (1983). Deficiency of an erythrocyte membrane protein with complement regulator activity in paroxysmal nocturnal hemoglobinuria. Proc. Natl. Acad. Sci. U.S.A. 80, 5430–5434. 74. Rosse, W. F., Logue, G. L., Adams, J., and Crookston, J. H. (1974). Mechanisms of immune lysis of the red cells in hereditary erythroblastic multinuclearity with a positive acidified serum test and paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 53, 31–43. 75. Holguin, M. H., Frederick, L. R., Bernshaw, N. J., Wilcox, L. A., and Parker, C. J. (1989). Isolation and characterization of a membrane protein from normal human erythrocytes that inhibits reactive lysis of the erythrocytes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 7–17. 76. Nose, M., Katoh, M., Okada, N., Kyogoku, M., and Okada, H. (1990). Tissue distribution of HRF20, a novel factor preventing the membrane attack of homologous complement, and its predominant expression on endothelial cells in vivo. Immunology 70, 145–149. 77. Sugita, Y., Nakano, Y., Oda, E., Noda, K., Tobe, T., Miura, N.-H., and Tomita, M. (1993). Determination of carboxyl-terminal residue and disulfide bonds of MACIF (CD59), a glycosylphosphatidylinositol-anchored membrane protein. J. Biochem. (Tokyo) 14, 473–477. 78. Meri, S., Morgan, B. P., Davies, A., Daniels, R. H., Olavesen, M. G., Waldmann, H., and Lachmann, P. J. (1990). Human protectin (CD59), an 18,000–20,000 MW complement lysis restricting factor, inhibits C5b-8 catalysed insertion of C9 into lipid bilayers. Immunology 71, 1–9. 79. Holguin, M. H., Wilcox, L. A., Bernshaw, N. J., Rosse, W. F., and Parker, C. J. (1989). Relationship between the membrane inhibitor of reactive lysis and the erythrocyte phenotypes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 1387–1394. 80. Sugarman, J., Devine, D. V., and Rosse, W. F. (1986). Structural and functional differences between decay-accelerating factor and red cell acetylcholinesterase. Blood 68, 680–684. 81. Low, M. G., and Finean, J. B. (1977). Release of alkaline phosphatase from membranes by a phosphatidylinositol-specific phospholipase C. Biochem. J. 167, 281–284. 82. Low, M. G. (1989). Glycosyl-phosphatidylinositol: a versatile anchor for cell surface proteins. FASEB 3, 1600–1608. 83. Davitz, M. A., Low, M. G., and Nussenzweig, V. (1986). Release of decay-accelerating factor (DAF) from the cell membrane by phosphatidylinositol specific phospholipase C (PIPLC). J. Exp. Med. 163, 1150–1161. 84. Holguin, M. H., Wilcox, L. A., Bernshaw, N. J., Rosse, W. F., and Parker, C. J. (1990). Erythrocyte membrane inhibitor of reactive lysis: Effects of phosphatidylinositol-specific phospholipase C on the isolated and cell-associated protein. Blood 75, 284–289. 85. Takahashi, M., Takeda, J., Hirose, S., Hyman, R., Inoue, N., Miyata, T., Ueda, E., Kitani, T., Medof, M. E., and Kinoshita, T. (1993). Deficient biosynthesis of N-acetylglucosaminylphosphatidylinositol, the first intermediate of glycosyl phosphatidylinositol anchor biosynthesis, in cell lines established from patients with paroxysmal nocturnal hemoglobinuria. J. Exp. Med. 177, 517–521. 86. Brubaker, L., Essig, L. J., and Mengel, C. E. (1977). Neutrophil life span in paroxysmal nocturnal hemoglobinuria. Blood 50, 657–662. 87. Dacie, J. V., and Lewis, S. M. (1961). Paroxysmal nocturnal haemoglobinuria: variation in clinical severity and association with bone marrow hypoplasia. Br. J. Haematol. 7, 442–457. 88. Lewis, S. M., and Dacie, J. V. (1967). The aplastic anaemia-paroxysmal nocturnal haemoglobinuria syndrome. Br. J. Haematol. 13, 236–251. 89. Nissen, C., Moser, Y., dalle Carbonare, V., Gratwohl, A., and Speck, B. (1989). Complete recovery of marrow function after treatment with anti-lymphocyte globulin is associated with high, whereas early failure and development of paroxysmal nocturnal haemoglobinuria are associated with low endogenous G-CSA-release. Br. J. Haematol 72, 573–577. 90. Tichelli, A., Gratwohl, A., Nissen, C., and Speck, B. (1994). Late clonal complications in severe aplastic anemia. Leuk. Lymphoma 12, 167–175.

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91. Dacie, J. V. (1963). Paroxysmal nocturnal haemoglobinuria. Proc. R. Soc. Med. 56, 587–596. 92. Cohen, P., Gardner, F. H., and Barnett, G. O. (1961). Reclassification of the thrombocytopenias by the Cr51-labeling method for measuring platelet life span. N. Eng. J. Med. 263, 1264–1268. 93. Devine, D. V., Siegel, R. S., and Rosse, W. F. (1987). Interactions of the platelets in paroxysmal nocturnal hemoglobinuria with complement. Relationship to defects in the regulation of complement and to platelet survival in vivo. J. Clin. Invest. 79, 131–137. 94. Sultan, C., Marquet, M., and Joffroy, Y. (1973). Etude de dysmyelopoieses acquises idiopathiques en culture de moelle in vitro. Nouv. Rev. Fr. Hematol. 13, 431–426. 95. Tumen, J., Kline, L. B., Fay, J. W., Scullin, D. C., Reisner, E. G., Rosse, W. F., and Huang, A. T. (1980). Complement sensitivity of paroxysmal nocturnal hemoglobinuria bone marrow cells. Blood 55, 1040–1046. 96. Dacie, J. V. (1999). Paroxysmal nocturnal haemoglobinuria. In ‘‘The Haemolytic Anaemias: Drug and Chemical Induced Haemolytic Anaemias, Paroxysmal Nocturnal Haemoglobinuria, and Haemolytic Disease of the Newborn,’’ pp. 139–330. Churchill Livingston, London.

2 Genetics of PNH Lucio Luzzatto and Khe´doudja Nafa Department of Human Genetics, Memorial Sloan Kettering Cancer Center, New York, New York 10021

Paroxysmal nocturnal hemoglobinuria (PNH) is a disorder encountered in all populations throughout the world. The disease can affect individuals of any age and of diverse socioeconomic groups. PNH has never been reported as a congenital disease and there is no family clustering. Thus, PNH is an acquired disease. Since the development of the acidified serum test [1, 2], it has been clear that intravascular hemolysis, which gives the disease its name, is due to an intrinsic abnormality of the red cell. In contrast, all other acquired hemolytic anemias are due to extracorpuscular causes. Therefore, this pathogenic combination— intracorpuscular red cell defect and acquired genetics—sets PNH apart from every hemolytic anemia. It has led to the formulation of PNH as a clonal disorder due to a somatic mutation [3]. Direct experimental evidence supporting this idea was obtained nearly three decades ago in PNH patients who were heterozygous for the X-linked gene encoding glucose-6-phosphate dehydrogenase [4] and has been fully confirmed since the cloning of the PIG-A gene [5].

Biochemical Genetics of PNH in Somatic Cells PNH belongs not to the study of inherited diseases but to somatic cell genetics. The biochemical abnormality of PNH cells was first identified by studying lymphoblastoid cell lines (LCL) that displayed the PNH phenotype, obtained through immortalization of peripheral blood lymphocytes from PNH patients. These were compared to LCLs with normal phenotype obtained from the same patients, and which therefore could be expected to be isogeneic except for the gene that must have undergone the somatic mutation that caused PNH. Comparative analysis of these two sets of cell lines revealed normal synthesis of phosphatidylinositol but failure to incorporate mannose or N-acetylglucosamine (GlcNAc), thus pinpointing a block at the level of the GlcNAc transfer [6, 7]. PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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The Dominance Test The pattern of inheritance of the PNH abnormality could not be determined by conventional family studies, because no kindred with inheritance of this condition has been available. However, a dominance test could be carried out using somatic cells. Human–human somatic cell hybrids were constructed by fusing PNH LCLs (see above) with normal cells. When hybrid clones were isolated and appropriately characterized, they had a normal phenotype [8]: thus, the PNH mutation is recessive with respect to the normal allele (Fig. 2-1). This immediately suggested that the PNH mutation entailed a loss rather than a gain of function.

Figure 2-1 The PNH phenotype is recessive in somatic cell hybrids (modified from [8] with permission from Kluwer-Plenum).

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2. Genetics of PNH

An intriguing paradox with respect to the somatic mutation model of the pathogenesis of PNH was also apparent: if the mutant gene were dominant, a somatic mutation would cause PNH; however, if the mutant gene were recessive, two mutations would be required for the PNH phenotype to manifest itself. The (almost) obligate inference was that the PNH gene must be X linked [8].

The PIG-A Gene The PIG-A cDNA was isolated in 1993 [5] by expression cloning, through its ability to restore the expression of glycosylphosphatidylinositol (GPI)-anchored proteins on the surface of the cells belonging to a mutant human lymphoid cell line (JY5), deficient in these proteins (the gene was called PIG-A for phosphatidyl inositol glycan complementation group A) (see Table 2-1 and Fig. 2-2). PIG-A was mapped to the short arm of the X chromosome by the use of somatic cell hybrids and by in situ hybridization [9], and subsequently finely mapped between DXS987 and DXS7169 on Xp22.1, within a region almost completely covered by physical contigs [10]. The human PIG-A gene is about 17 kb in size and consists of six exons [9, 11, 12], while the mRNA contains 3589 nt. The coding region is 1452 nt long; exon 1 (23 nt) and 62 nt of exon 2 are noncoding. In addition to the major mRNA species, two minor species of 3.3 and 2.8 kb, respectively, have been observed on Northern analysis; they arise from two alternative splice sites at nt position 58 and 342 (in exon 2); from sequence

TABLE 2-1 Characteristics of the PIG-A Gene Genomic Location Subject to X inactivation Size of the gene (kb) Number of exons Coding exons cDNA (bp) Isolated by expression cloning in 1993 Size in nucleotides 5⬘ untranslated region 3⬘ untranslated region Coding region Protein (amino acids) CpG dinucleotides Pseudogene Polymorphisms In exons In introns a

Between DX987 and DX7169 [10].

Xp22.1a Yes 17 6 5

3589 85 2049 1452 484 23 12q21 1 (55C씮T) None known

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Figure 2-3 Multiple mRNA species from the PIG-A gene. This picture is an autoradiograph of RTPCR analysis of PIG-A mRNA from PNH cell lines. Three major amplification products are detected. The largest amplification product corresponds to the coding region, and the two additional products are due to the presence of alternative splicing sites (see Fig. 2-4). In the HH8/3P and HH5/7P cell lines, frameshift mutations were identified (336GC씮T and 634insTAGAT in exon 2 of the PIG-A gene, respectively). In the HH20/9P and HH16/7P cell lines, the patterns are indistinguishable from that of the normal JY cell line. Transfection of the 1500-bp product of HH20/9P and HH16/7P failed to restore CD59 expression. A 464 C-to-T base change and a 1348 AT씮C were identified in each of the cell lines HH16/7P and HH20/9P, respectively. (Reproduced from [16], with permission from Oxford University Press.)

analysis it can be inferred that they are nonfunctional (see Figs. 2-3 and 2-4) [12, 13]. The PIG-A gene promoter has four CAAT boxes, two AP motifs, and one CRE motif (Fig. 2-5). There is no TATA box (12). A nonfunctional PIG-A pseudogene has been mapped to chromosome 12q21 [9].

The PIG-A Protein The predicted protein product of PIG-A consists of 484 amino acids [5]. A hydrophobic region near the carboxyl terminus may be a transmembrane domain (amino acids 415–442). The hydrophilic carboxyl terminal region of 42 residues corresponds to the luminal domain of PIG-A [14]. There is no amino-terminal hydrophobic signal sequence. A region of 92 amino acids spanning residues 304–395 is homologous (27% identity) to the bacterial glucosaminyl N-acetyl transferase, Rfa K, which is involved in the synthesis of liposaccharides. By sitedirected mutagenesis, only the first 23 luminal residues that immediately follow the transmembrane domain appear essential for the function of PIG-A. Watanabe et al. [15], have also shown that PIG-A, PIG-H, PIG-C, and GPI1 are subunits of a protein complex that mediates the first reaction step of the GPI anchor biosynthesis, the transfer of GlcNAc unto phosphatidylinositol. This model is consistent with the biochemical studies described above. That PIG-A is part of an enzyme complex is also compatible with the recessive character of PNH mutations. The PIG-A mutations that cause PNH are those that produce partial or total loss of the enzymatic function of the PIG-A protein.

Figure 2-4 Splicing pattern and splicing mutations in the PIG-A transcript. In the processing of the normal PIG-A gene, three types of exon–exon junctions take place. Mutations in types 1 and 2 exon–exon junctions cause frameshift mutations; mutations in type 3 lead to exon skipping (numbers 1, 2, and 3 correspond to the first, second, and third nucleotides of the codon). The two cryptic splice sites in the normal PIG-A gene are shown (arrows). (A) The 5⬘ splice site mutation in intron 3 (IVS3⫹1g씮a) leads to the use of the alternative splice (AGGTAC) in exon 2 at position 342-343 and the deletion of 374 bp of exon 2 and the entire exon 3 (putative PIG-A protein of 315 aa) [39]. (B) Mutations either in the 3⬘ of intron 3 [acceptor splice site] or in the 5⬘ of the intron 4 [donors splice site] cause skipping of exon 4 with production of an out-of-frame mature mRNA introducing an early stop codon. (C) The IVS3-1g/GGT씮IVS3-1a/GGT mutation in the genomic DNA causes loss of the first nucleotide of exon 4 (849G) in the mRNA and a premature chain termination at position 290 without the deletion of exon 4 [40]. (D) The IVS5⫹lg씮a and IVS5⫹2delt mutations in intron 5 result in the loss of exon 5 and a truncated PIG-A protein of only 415 aa [11, 26, 27, 45].

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Figure 2-5 Structure of the PIG-A promoter. The four CAAT boxes, the two AP-2 motifs, and one CRE motif are shown in bold type. Region corresponding to the deletion of 737 bp in the promoter (Table 2-3, mutation number 3) is underlined [40]. The A of the initiation codon is the nucleotide position 1 of the translated region.

2. Genetics of PNH

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Detecting Mutations in the PIG-A Gene In principle, the PNH phenotype could arise from a metabolic block at any step in the GPI anchor biosynthesis. However, there are several good reasons to presume that PIG-A is mutated in most and perhaps all patients with PNH. (i) The PIG-A cDNA was shown to correct in vitro the PNH phenotype in all four of the first lymphoblastoid cell lines tested, which had been obtained from four different patients [16]. (ii) PIG-A mutations are present in about 80% of patients with PNH that have been appropriately investigated (see below). (iii) All of the other genes in the GPI biosynthetic pathway that have been mapped to date are autosomal [17–22]; their functional inactivation would almost certainly require that both alleles be mutated, a very unlikely event at the somatic cell level (although it may be the case in a unique family with PNH, possibly the single exception to the rule that PNH is an acquired disorder [23]). In view of these considerations, genetic testing for PNH consists in practice in searching for a mutation in PIG-A. In this chapter, we will concentrate on the approaches that have been used more extensively and particularly on the methodology that we have found most effective in our experience.

Nature of Sample Because a PIG-A mutation giving rise to PNH must have occurred in a multipotent hematopoietic stem cell [1], in theory the analysis can be performed in any derived cell: unfractionated peripheral blood white cells [24], individual types of white cells [25–33], bone marrow cells [11, 34], B cell lines, and T cell lines [35–41]. Because the proportion of granulocytes belonging to the PNH clone is usually larger than that of any other cell type, this has been our preferred material in most cases. The technique we follow has been fully described recently [42, 43]. Archival material, such as stained or unstained bone marrow slides, can be also used [44].

Genomic DNA vs. cDNA PIG-A mutations in the coding region, which are the majority, can be identified by cDNA amplification (in this case, peripheral blood reticulocytes are an additional tissue source). However, obtaining good quality RNA from which to make cDNA is in general more difficult than direct amplification of genomic DNA, especially if the sample has been stored or shipped. In addition, this approach will miss some of the splice sites mutations. Therefore, we normally use genomic DNA.

Mutation Analysis Our algorithm (see Fig. 2-6) is based on PCR amplification of the entire coding region in four fragments, (either by RT-PCR or by PCR on a genomic DNA template), followed by the screening for the presence of abnormal fragments by

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Figure 2-6 Strategy for identifying mutations in the PIG-A gene. The numbers shown refer to a series of 49 patients reported by Nafa et al. (1998) from which this figure is reproduced [43].

heteroduplex analysis (HA) and single strand conformation analysis (SSCA), followed by nucleotide sequencing to identify the mutation. For HA and SSCA, the amplification products are digested with different restriction enzyme mixtures [16, 35, 36]. Conditions favoring heteroduplex formation prevail during PCR cycling, thanks to the coexistence in PNH patients of the normal with the abnormal PIG-A sequence. If the abnormal DNA fragment is abundant, direct sequencing can be performed; otherwise, subcloning into phage M13 is required (see Figs. 2-7 and 2-8). In order to safeguard against misinterpreting PCR artifacts, single base changes should be considered responsible for PNH only if found in at least two out of several individual M13 clones. With a small deletion/insertion, a single abnormal result in only one M13 clone can be more confidently interpreted, because Taq polymerase is unlikely to produce such errors although it is still safer to sequence additional M13 clones. If the mutation causes a change in a restriction site, its presence can be conveniently confirmed by the appropriate digestion of PCR-amplified uncloned

2. Genetics of PNH

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Figure 2-7 Example of PIG-A mutations revealed by single strand conformation analysis. (A) In MSK11, a male patient with 93% PNH PMN, an abnormal band is seen in each of three digests of amplified exon 2 fragment. (B) Restriction map of exon 2, as amplified for SSCA analysis, showing the restriction fragments for which abnormal bands were observed (in bold). The combined data narrow down the location of the mutation to a 71-bp region between the TaqI site at position nt 228 and the AccI site at position nt 299 (arrows). (C) Direct nucleotide sequencing of exon 2 indeed revealed a deletion of cytidine at nt position 259 resulting in a frameshift and premature chain termination at codon 93. This mutation eliminates a HaeIII restriction site. (D) Confirmation of the mutation by restriction enzyme digestion of the PCR product of exon 2.

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Figure 2-8 Example of two PNH clones with two different mutations in exon 2 of the PIG-A gene in patient HH60. (Top) Four M13 clones have a 143G씮A substitution (Nos. 1, 5, 6, and 8). (Bottom) Two M13 clones have a 715G씮A substitution (Nos. 4 and 7). None of the M13 clones have both mutations. The remaining M13 clones (Nos. 2 and 3) are normal. (Reprinted from [43].)

DNA. To confirm that the observed changes have arisen through somatic mutations, DNA from the patient’s mononuclear cells (which usually have a much lower proportion of PNH cells than granulocytes) can serve as an internal control. Southern blotting analysis is not routinely carried out because large deletions within the PIG-A gene appear to be rare, although, they can be encountered (see Figs. 2-2 and 2-9). A deletion of the entire PIG-A gene would be missed in a female patient, due to the presence of an intact gene on the inactive X chromosome; such a situation would be suspected by quantification of the hybridization signal normalized to an internal control, like the PIG-A pseudogene. PIG-A mutations causing premature termination of translation, which are the majority, could be detected by the so-called protein truncation test (PTT) [45] (see Fig. 2-10). Finally, a more laborious but elegant approach is a functional assay, whereby the cDNA of the entire coding region from a patient with PNH is cloned into a mammalian expression vector and transfected into a GPI-deficient cell line, for instance, JY5 [11, 16, 26, 28, 34, 38]. The cDNA from clones that fail to rescue the PNH phenotype are then sequenced. In principle, this is the only technique to prove that a missense mutation is functionally relevant; however, even this approach may not be conclusive, because transfectants may contain multiple gene copies, and therefore the level of expression may not mimic that prevailing in vivo from a single copy of the mutant gene.

Figure 2-9 A rare deletion of the PIG-A gene. (A) When the DNA is digested with Bgl II and hybridized with a full-length PIG-A cDNA probe, there are four restriction fragments, three corresponding to the PIG-A gene and one to the ⌿PIG-A gene. In MSK8, the ⌿PIG-A gene band is still present but the other three are missing. We infer that the entire PIG-A gene is deleted in this patient. To further characterize the extent of the deletion, a filter containing EcoRI-digested DNA was hybridized with the FIGF (c-fos-induced growth factor) cDNA, which also maps to Xp22.1 [47]. The deletion extended as far as the FIGF gene and two other microsatellite markers (DXS207 and DXS1053) (unpublished data, not shown). (B) Diagram of the Xp22.1 region. The FIGF and PIG-A genes lie next to each other in a head-to-tail orientation separated by only 12 kb. The FIGF spans approximately 50 kb. Therefore, the deleted region in MSK8 is more than 62 kb (bold line).

Figure 2-10 A splicing mutation in the PIG-A gene revealed by the protein truncation test (PTT). The RT-PCR products of PIG-A mRNA from the PNH patient and normal controls were analyzed by the in vitro transcription/translation test. A major band corresponding to a shorter polypeptide (calculated size 46.7 kD) was present in the patient sample (lane 4), in addition to a very faint normal size polypeptide (size 53.3kD). Lane 1, molecular size marker; lanes 2 and 3, translation products from controls. (Reproduced from [45] with permission from Blackwell Science Ltd.)

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Figure 2-11 AvaI polymorphism in exon 2 of the PIG-A gene in HH55 family. The X chromosome with the loss of AvaI site is inherited from the father (subject I1).

Germ Line PIG-A Mutations Thus far, only one instance of an inherited mutation of the PIG-A gene has been reported: a single nucleotide substitution (C55T) in exon 2, which causes a replacement of tryptophan for arginine at aa position 19 (easy to detect because it causes the loss of an AvaI restriction site at position nt 55) (see Fig. 2-11). The 55T PIG-A allele is polymorphic, with an estimated heterozygote frequency of about 4.8% in PNH patients (Table 2-2). This mutation does not cause PNH, because constitutional loss of the AvaI site has been observed in subjects not suffering from PNH. However, it is interesting that we and others first observed this mutation in patients with PNH: to date, in the heterozygous state in five (out of 106) patients and in the homozygous state in one [39, 42, 43, 46]. A thymidine to cytidine substitution at position ⫺23 in intron 3 was found in one PNH patient, and a silent mutation in exon 4 reported in another [43, 26]. Although these mutations were constitutional (as they were found also in the non-PNH cells of the respective patients), they have not been formally proven to be inherited.

TABLE 2-2 Frequency of the 55C씮 씮 T, 19 Arg씮 씮 Trp Polymorphism Number

PHN Algerians (Non-PNH)

Male

Female

Hemizygote 55 T

Heterozygote 55 C/T

Allelic frequency

26 43

39 5

0 1

5 0

4.8% (5/104) 1.9% (1/53)

2. Genetics of PNH

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The Spectrum of Somatic Mutations in the PIG-A Gene To date, a total of 174 somatic mutations in the PIG-A gene have been identified by different investigators in more than 28 reports in 146 patients with PNH (Table 2-3). Among them, 158 were unique mutations and 16 were recurrent mutations (reported in more than one patient). Among the 174 somatic mutations, 135 mutations are such that we can predict complete functional inactivation of the PIG-A gene product (PIG-A⬚): 3 large deletions, 96 frameshift mutations, 19 nonsense mutations, and 17 splice site mutations. Thirty-five missense mutations and four in-frame small deletions have also been described.

Large Deletions This type of mutation seems to be relatively rare, as only three instances have been reported (Table 2-3, lines 1–3). The first case (HH22) was a 4-kb deletion encompassing exons 3, 4, and 5 [16]. In the second (MSK8), the entire PIG-A gene was deleted [43]; the size of the deletion has not been established, but it must be more than 62 kb ([47] and unpublished data). The third instance was a 737-bp deletion, removing 137 nt of the promoter, exon 1 (23 nt) and 577 nt of intron 1 (see Figs. 2-2 and 2-5) [40].

Frameshift Mutations There are 96 mutations of this type in the database: 63 are small deletions of 1 to 19 nt; 22 are small insertions of 1 to 19 nt; 7 are deletions/insertions, and 4 are duplications of 7 to 88 nt [43, 48].

Splice Site Mutations Among the 17 splicing mutations, 15 consist of single base pair substitutions and 2 single nucleotide deletions [11, 37]. About one-third of the splice site mutations are recurrent (see Tables 2-3 and 2-4 and Fig. 2-4), including the out-of-frame deletion of exon 4 (four times) and the in-frame deletion of exon 5 (exon skipping, three times).

Point Mutations Nineteen nonsense mutations and 35 missense mutations have been published. The nonsense mutations are spread throughout the entire coding region, while the missense mutations are largely confined to exon 2 of the PIG-A gene. Four small in-frame deletions are also localized in exon 2: they consist of the deletion of 3 nt (two cases), 9 nt, and 24 nt [27, 33, 43, 52]. In four patients (see Table 2-3, lines 9, 119, 151, and 180), the changes were very close to each other, presumably resulting from a single mutational event that occurred in an early hematopoietic stem cell [37, 43, 49].

34 TABLE 2-3 Somatic Mutations of the PIG-A Gene Mutation number

Exon/intron

Nucleotide number

Base change

Codon number

aa change

Consequence

Detection method

Reference

1 2 3 4 5

Entire gene del Exon 3,4,5 Promo-intron 1 Exon 2 Exon 2

— — — 16 39

Large deletion Large deletion Del 737 bp G씮A Del C

— — — 6 13

— Stop 294 — gly씮stop Stop 60

Complete loss Frameshift — Nonsense Frameshift

Southern Southern Southern Sequencing RT-PCR

Nafa et al., 1998 [43] Bessler et al., 1994 [36] Endo et al., 1996 [40] Nafa et al., 1998 [44] Nagarajan et al., 1995 [39]

6

Exon 2

55

C씮T

19

arg씮trp

Missense

HA, SSCA

Araten et al., 1999 [53]

7

Exon 2

55

C씮T

19

arg씮trp

Missense

HA, SSCA

Araten et al., 1999 [53]

8 9

Exon 2 Exon 2

68 102–112/98

Del G Del 11 nt/A씮G

23 34/33

Stop 60 Stop 36/his씮arg

Sequencing RT-PCR

Lin et al. 1997 [24] Ostendorf et al., 1995 [37]

10 11 12 13 14 15 16 17 18

Exon Exon Exon Exon Exon Exon Exon Exon Exon

2 2 2 2 2 2 2 2 2

104 106 118 118 142 143 143 145 161

T씮A Ins AT Del G G씮C G씮A G씮A G씮C GT씮AA Ins T

35 36 40 40 48 48 48 49 54

ile씮lys Stop 61 Stop 60 asp씮his gly씮ser gly씮asp gly씮ala val씮leu Stop 61

Frameshift Frameshift/ missense Missense Frameshift Frameshift Missense Missense Missense Missense Missense Frameshift

RT-PCR, HA RT-PCR HA, SSCA HA HA HA, SSCA SSCA HA, cDNA HA

Yamada et al., 1995 [26] Endo et al., 1996 [41] Nafa et al., 1995 [42] Nafa et al., 1998 [43] Nafa et al., 1995 [42] Nafa et al., 1998 [43] Nafa et al., 1998 [43] Pramoonjago et al., 1999 [29] Nafa et al., 1998 [43]

19 20

Exon 2 Exon 2

163 163

C씮T C씮T

55 55

gln씮stop gln씮stop

Nonsense Nonsense

F, SSCA RT-PCR

Bessler et al., 1994 [35] Nagarajan et al., 1995 [39]

21 22 23 24 25 26 27 28 29 30

Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon

166 167 172 183 189 192 193 196 211 211/251

Del CT T씮C C씮T Del T Del A Del G Del C Ins AT A씮G A씮G/C씮T

56 56 58 61 63 64 65 66 71 71/84

Stop 61 leu씮pro gln씮stop Stop 68 Stop 68 Stop 68 Stop 68 Stop 69 thr씮ala thr씮ala/thr씮ile

Frameshift Missense Nonsense Frameshift Frameshift Frameshift Frameshift Frameshift Missense Missense

HA, SSCA HA, SSCA SSCA, seq HA, SSCA HA, SSCA HA, SSCA HA, cDNA HA, SSCA Sequencing SSCA

Nafa et al., 1998 [43] Araten et al., 1999 [53] Merk et al., 1997 [52] Bessler et al., 1994 [36] Nafa et al., 1995 [42] Nafa et al., 1998 [43] Pramoonjago et al., 1999 [29] Araten et al., 1999 [53] Nafa et al., 1998 [44] Nafa et al., 1998 [44]

2 2 2 2 2 2 2 2 2 2

Comments

B-LCL T cell clone

Healthy donor, AvaI(-) Healthy donor, AvaI(-)

T cell clone

B-LCL; AvaI(-)

AvaI(-) Healthy donor B-LCL; AvaI(-)

Healthy donor

31 32

Exon 2 Exon 2

229 229

C씮T C씮T

77 77

arg씮stop arg씮stop

Nonsense Nonsense

HA, SSCA

Araten et al., 1999 [53] Pramoonjago et al., 1999 [29]

33 34 35 36 37 38 39 40 41 42 43

Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon

242 246 247 250 250 259 265–288 269 275 289 290

Del G Ins A Del C(CTC씮TC) Del A Ins GT Del C Del 24 nt Ins T Ins CTATTACT A씮T Ins A

81 82 83 84 83 84 89–96 90 92 97 97

Stop 94 Stop 82 Stop 94 Stop 94 Stop 95 Stop 94 Del of 8 aa Stop 129 Stop 97 met씮leu Stop 129

Frameshift Frameshift Frameshift Frameshift Frameshift Frameshift In frame (476) Frameshift Frameshift Missense Frameshift

HA RT-PCR, HA HA, cDNA SSCA RT-PCR HA, SSCA SSCA RT-PCR, HA HA SSCA HA

Nafa et al., 1998 [43] Miyata et al., 1994 [38] Pramoonjago et al., 1999 [29] Merk et al., 1997 [52] Ware et al., 1994 [25] Nafa et al., 1998 [43] Merk et al., 1997 [52] Yamada et al., 1995 [26] Azenishi et al., 1999 [33] Merk et al., 1997 [52] Azenishi et al., 1999 [33]

44 45

Exon 2 Exon 2

294 294

C씮A C씮A

98 98

tyr씮stop tyr씮stop

Nonsense Nonsense

RNA-SSCA HA, SSCA

Savoia et al., 1996 [30] Araten et al., 1999 [53]

46 47

Exon 2 Exon 2

298 298

C씮T C씮T

100 100

gln씮stop gln씮stop

Nonsense Nonsense

RT-PCR, HA SSCA

Yamada et al., 1995 [26] Nafa et al., 1995 [42]

48 49

Exon 2 Exon 2

304 317

A씮T T씮A

102 106

thr씮pro leu씮his

Missense Missense

RT-PCR SSCA, HA

Endo et al., 1996 [40] Du et al., 1997 [32]

T cell clone

50 51

Exon 2 Exon 2

336 336

GC씮T GC씮T

112–113 112–113

Stop 124 Stop 124

Frameshift Frameshift

HA, SSCA RT-PCR, HA

Bessler et al., 1994 [16] Yamada et al., 1995 [26]

B-LCL

52 53

Exon 2 Exon 2

338 338

T씮C T씮C

113 113

leu씮pro leu씮pro

Missense Missense

HA, SSCA SSCA

Nishimura et al., 1997 [34] Merk et al., 1997 [52]

B-LCL

54 55

Exon 2 Exon 2

356–357 362

Del GG Del GAG

119 121–122

Stop 128 Del glu

Frameshift In frame

HA, cDNA HA, SSCA

Pramoonjago et al., 1999 [29] Nafa et al., 1998 [43]

56 57 58 59

Exon Exon Exon Exon

383 383 383 383

A씮G A씮G A씮G A씮G

128 128 128 128

his씮arg his씮arg his씮arg his씮arg

Missense Missense Missense Missense

HA-SSCA RT-PCR, FA HA, cDNA RT-PCR, FA

Nafa et al., 1998 [43] Yamada et al., 1995 [26] Pramoonjago et al., 1999 [29] Yamada et al., 1995 [26]

60 61 62

Exon 2 Exon 2 Exon 2

384 387 394/86

TAGTTC씮A T씮A Del T/G씮C

128 129 132/29

Stop 135 ser씮arg Stop 171/arg씮thr

HA, SSCA HA, SSCA HA

Nafa et al., 1995 [42] Nafa et al., 1995 [42] Pakdeesuwan et al., 1997 [28]

63 64 65 66

Exon Exon Exon Exon

404 408 431 443

Ins T Del T Del C AGA씮T

135 136 144 148

Stop Stop Stop Stop

Frameshift Missense Frameshift/ missense Frameshift Frameshift Frameshift Frameshift

RT-PCR, HA RT-PCR, HA RT-PCR HA

Pramoonjago et al., 1995 [22] Miyata et al., 1994 [38] Ware et al., 1994 [25] Nafa et al., 1998 [43]

2 2 2 2 2 2 2 2 2 2 2

2 2 2 2

2 2 2 2

161 173 198 160

Healthy donor

Healthy donor

MDS/PNH

35

continues

36 TABLE 2-3 Mutation number

continued

Exon/intron

Nucleotide number

Base change

Codon number

aa change

Consequence

Detection method

Reference

67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84

Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon

2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2

448 449 451–453 460 464 467 471 491 503 506 520 548 549 556 564 568 571 575

G씮A T씮C Del TTC Ins A C씮T T씮G Del T C씮A A씮G Ins A Ins GT G씮T T씮G Del T Del T G씮T Del A Del C

150 150 151 154 155 156 157 164 168 169 174 183 183 186 188 190 191 192

val씮ile val씮ala 483 Stop 160 ser씮phe leu씮arg Stop 171 ser씮stop asn씮ser Stop 177 Stop 195 cys씮phe cys씮trp Stop 194 Stop 194 glu씮stop Stop 194 Stop 194

Missense Missense In frame Frameshift Missense Missense Frameshift Nonsense Missense Frameshift Frameshift Missense Missense Frameshift Frameshift Nonsense Frameshift Frameshift

SSCA, HA SSCA, HA RT-PCR, HA RNA-SSCA RT-PCR SSCA SSCA, HA HA SSCA HA, cDNA HA SSCA Sequencing HA, SSCA RT-PCR SSCA FA, RT-PCR RT-PCR

Du et al., 1997 [32] Du et al., 1997 [32] Pramoonjago et al., 1995 [22] Savoia et al., 1996 [30] Bessler et al., 1994 [16] Bessler et al., 1994 [35] Du et al., 1997 [32] Pakdeesuwan et al., 1997 [28] Merk et al., 1996 [52] Pramoonjago et al., 1999 [29] Nafa et al., 1995 [42] Nafa et al., 1995 [42] Rollinson et al., 1997 [54] Nafa et al., 1998 [43] Endo et al., 1996 [40] Merk et al., 1997 [52] Yamada et al., 1995 [26] Nagarajan et al., 1995 [39]

85 86 87 88

Exon Exon Exon Exon

2 2 2 2

577 577 577 577

Del Del Del Del

193 193 193 193

Stop Stop Stop Stop

Frameshift Frameshift Frameshift Frameshift

RT-PCR, RT-PCR, RT-PCR, RT-PCR,

HA HA HA HA

Merk et al., 1997 [52] Merk et al., 1997 [52] Merk et al., 1997 [52] Pramoonjago et al., 1995 [22]

89 90 91 92 93 94 95 96 97 98 99

Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon

2 2 2 2 2 2 2 2 2 2 2

578 609 614 627 633 634 634 635 663 668 678–686

Ins ATGT Del GT T씮A Del T Del T Ins TAGAT Del C Del C A씮C Del ATGA Del 9 nt (AACTATTGT)

193 203 205 209 211 211 212 211 221 223 226–229

Stop 202 Stop 207 val씮asp Stop 210 Stop 226 Stop 212 Stop 226 Stop 226 arg씮ser Stop 225 del 3 aa

Frameshift Frameshift Missense Frameshift Frameshift Frameshift Frameshift Frameshift Missense Frameshift In frame del

RT-PCR RT-PCR, HA HA, SSCA RT-PCR, HA HA, cDNA RT-PCR, HA HA, cDNA HA SSCA H-SSCP HA

Nagarajan et al., 1995 [39] Pramoonjago et al., 1995 [22] Araten et al., 1999 [53] Pramoonjago et al., 1995 [22] Pramoonjago et al., 1999 [29] Bessler et al., 1994 [16] Pramoonjago et al., 1999 [29] Azenishi et al., 1999 [33] Merk et al., 1997 [52] Nafa et al., 1998 [43] Azenishi et al., 1999 [33]

GTACT GTACT GTACT GTACT

199 199 199 199

Comments

B-LCL B-LCL

T cell clone NHL

NHL AA NHL

Healthy donor

B-LCL

100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119

Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon Exon

120 121 122

2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 3 3 3 3 3

696 680 692 693 715 715 716 717 743 753 762 767 770 773–791 782 787 803 816 820 824/829

Ins 19 nt (678–696) Del C T씮C Dup 88 nt Del G G씮A G씮T Del G Del C Del T T씮A Del A Del T Del 19 nt Ins A G씮A Del AG Del A Del G G씮C/Ins A

227–233 227 231 231 239 239 239 239 248 251 254 256 257 258 261 263 268 272 274 275/277

Stop 250 Stop 246 val씮ala Stop 237 Stop 246 gly씮arg gly씮val Stop 246 Stop 257 Stop 257 tyr씮stop Stop 258 Stop 257 Stop 284 Stop 282 gly씮arg Stop 280 Stop 290 Stop 289 arg씮pro/stop 282

HA, cDNA HA, SSCA SSCA RT-PCR HA SSCA HA RT-PCR RT-PCR, HA HA RT-PCR RT-PCR HA, SSCA HA HA HA, cDNA HA RT-PCR, HA SSCA HA, SSCA

Pramoonjago et al., 1999 [29] Nafa et al., 1995 [42] Merk et al., 1997 [52] Pavlu et al., 1997 [48] Nishimura et al., 1996 [31] Nafa et al., 1998 [43] Pakdeessuwan et al. 1997 [28] Ostendorf et al., 1995 [37] Pramoonjago et al., 1995 [22] Nafa et al., unpublished Nagarajan et al., 1995 [39] Nagarajan et al., 1995 [39] Nafa et al., 1995 [42] Nafa et al., 1998 [43] Nafa et al., 1995 [42] Pramoonjago et al.,1999 [29] Nafa et al., 1998 [43] Pramoonjago et al., 1995 [22] Nafa et al., 1995 [42] Nafa et al., 1998 [43]

Stop 293 284 Stop 290

Frameshift Frameshift Missense Frameshift Frameshift Missense Missense Frameshift Frameshift Frameshift Nonsense Frameshift Frameshift Frameshift Frameshift Missense Frameshift Frameshift Frameshift Missense/ frameshift Frameshift Del exon 4 Frameshift

Intron 3 Intron 3 Intron 3

⫹1 ⫺1 ⫺1

Del 507 nt — —

123 124 125 126 127

Intron 3 Exon 4 Exon 4 Exon 4 Exon 4

⫺2 851–860 859–960 860 874

128 129 130 131

Exon Exon Exon Exon

4 4 4 4

889 897 901 934

gt씮at ag/G씮--/G (del ag) ag/G씮aa/G; cDNA 849 Del G ag/G씮gg/G Del 10 nt Del TT Ins 2 nt (TTG씮TTTGG) Ins CAG-dup 25 nt (849–874) A씮G Ins T C씮T Del A

RT-PCR RT-PCR RT-PCR

Nagarajan et al., 1995 [39] Ostendorf et al., 1995 [37] Endo et al., 1996 [40]

— 284–287 287 287 292

— Stop 287 Stop 295 — Not determined

Frameshift Frameshift Frameshift Frameshift Frameshift

RT-PCR RT-PCR HA HA, cDNA HA-SSCA

Yamada et al., 1995 [26] Pramoonjago et al., 1995 [22] Azenishi et al., 1999 [33] Pramoonjago et al., 1999 [29] Nafa et al., 1998 [43]

297 299 301 312

asn씮asp Stop 312 gln씮stop Stop 329

Missense Frameshift Nonsense Frameshift

RT-PCR HA HA RT-PCR, HA

Ware et al., 1994 [25] Pramoonjago et al., 1995 [2] Nafa et al., 1998 [43] Yamada et al., 1995 [26]

132

Exon 4

967

Del A

323

Stop 329

Frameshift

HA, cDNA

Pramoonjago et al., 1999 [29]

133 134

Intron 4 Intron 4

⫹1 ⫹1

G/gt씮G/at G/gt씮G/at

— —

— —

Del exon 4 Del exon 4

SSCA SSCA

Nafa et al., 1995 [42] Yamada et al., 1995 [26]

Functional assay

AvaI(-)

B-LCL T cell clone, genomic DNA

cDNA not analyzed

Has neutral inherited mutation

continues

37

38 TABLE 2-3 Mutation number

continued

Exon/intron

Nucleotide number

Base change

Codon number

aa change

Consequence

135

Intron 4

⫹1

G/gt씮G/at

136

Intron 4

⫹1

G/gt씮G/at

—

—

137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157

Intron 4 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5 Exon 5

⫹1 983 985–986 986 987 1000 1003 1027 1032 1057 1058 1060 1099 1100 1110/1130 1111 1114 1132 1135 1141–1154 1160

G/gt씮G/ct Del T Del GT Ins T Ins T G씮A G씮T Del AA Del T A씮T Dup nt 1052–1058 Ins A A씮T AGT씮TG A씮G/T씮A Del T Del C Del C Ins T Del 14 nt G씮A

— 328 329 329 329 334 335 343 344 353 353 354 367 367/368 370/377 371 371 378 379 381 387

— Stop 329 Stop 337 Stop 338 Stop 338 gly씮ser gly씮stop Stop 348 Stop 352 lys씮stop Stop 358 Stop 357 lys씮stop Stop 380 thr씮thr/ile씮asn Stop 380 Stop 380 Stop 380 Stop 380 Stop 390 trp씮stop

158 159 160

Intron 5 Intron 5 Intron 5

⫹1 ⫹1 ⫹1

G/gt씮G/at G/gt씮G/at G/gt씮G/at

— — —

— — —

Del exon 5 Del exon 5 Del exon 5

—

—

Del exon 4

Detection method

Reference

Comments

SSCA

Nafa et al., 1998 [43]

cDNA not analyzed

Del exon 4

SSCA

Pramoonjago et al., 1995 [22]

Del exon 4 Frameshift Frameshift Frameshift Frameshift Missense Nonsense Frameshift Frameshift Nonsense Frameshift Frameshift Nonsense Frameshift Silent/missense Frameshift Frameshift Frameshift Frameshift Frameshift Nonsense

RT-PCR RT-PCR HA, seq HA HA, SSCA Sequencing HA, SSCA HA, SSCA HA, cDNA SSCA HA HA SSCA HA, cDNA RT-PCR HA, cDNA RNA-SSCA HA, SSCA HA RT-PCR RT-PCR

Ostendorf et al., 1995 [37] Ostendorf et al., 1995 [37] Azenishi et al., 1999 [33] Nafa et al., 1998 [43] Nishimura et al., 1997 [34] Lin et al., 1997 [24] Nishimura et al., 1997 [34] Nishimura et al., 1997 [34] Pramoonjago et al., 1999 [29] Merk et al., 1997 [52] Nafa et al., unpublished Nafa et al., 1995 [42] Nafa et al., 1998 [43] Pramoonjago et al., 1999 [29] Stafford et al., 1995 [49] Pramoonjago et al., 1999 [29] Savooia et al., 1996 [30] Nafa et al., 1995 [42] Azenishi et al., 1999 [33] Nagarajan et al., 1995 [39] Endo et al., 1996 [41]

T cell clone

RT-PCR RT-PCR RT-PCR/PTT

Yamada et al., 1995 [26] Pramoonjago et al., 1995 [22] Maugard et al., 1997 [45]

Exon skipping Exon skipping Exon skipping

B-LCL BFU-E/CFU mix BFU-E/CFU mix NHL

B-LCL

161

Intron 5

⫹2

G/gt씮G/g- (del t)

—

—

Splice site

—

Takeda et al., 1993 [11]

162 163

Intron 5 Intron 5

⫺2 ⫺2

ag씮gg ag씮gg

— —

— —

Splice site Splice site

HA, SSCA HA, cDNA

Nafa et al., 1998 [43] Pramoonjago et al., 1999 [29]

164 165

Intron 5 Intron 5

⫺1 ⫺1

G씮A G씮A

— —

— —

Splice site Splice site

HA, cDNA HA, cDNA

Pramoonjago et al., 1999 [29] Pramoonjago et al., 1999 [29]

166

Exon 6

1195–1204

399–402

Stop 420

Frameshift

HA

Azenishi et al., 1999 [33]

167 168 169


1214 1220 1220

Del 10 nt (GACCGGGTAT) C씮G Del 7 nt Del T

405 407 407

ala씮val Stop 421 Stop 442

Missense Frameshift Frameshift

Sequencing SSCA, HA RT-PCR, HA

Lin et al., 1997 [24] Nafa et al., 1998 [43] Pramoonjago et al., 1995 [22]

170 171

Exon 6 Exon 6

1234 1234

Del C Del C

412 412

Stop 423 Stop 423


RT-PCR, HA SSCA

Pramoonjago et al., 1995 [22] Nafa et al., 1998 [43]

172 173 174 175

Exon Exon Exon Exon

1254–1264 1260 1280 1291

Del 11 nt/ins G C씮A Del T T씮A

418–422 420 427 431

Stop 420 cys씮stop Stop 442 leu씮stop

Frameshift Nonsense Frameshift Nonsense

HA, cDNA HA, SSCA RT-PCR, HA RT-PCR, FA

Pramoonjago et al., 1999 [29] Nafa et al., 1998 [43] Pramoonjago et al., 1995 [22] Yamada et al., 1995 [26]

TM (nt 1246–1326) TM (nt 1246–1326) TM (nt 1246–1326)

176 177

Exon 6 Exon 6

1309 1309

Del C Del C

437 437

Stop 442 Stop 442


RT-PCR, HA HA-SSCA

Yamada et al., 1995 [26] Bessler et al., 1994 [36]

TM (nt 1246–1326) TM, B-LCL

178 179 180


1323 1331 1331/1335

Del CT G씮A G씮A/G씮A

441–442 444 444/445

Stop 451 trp씮stop trp씮stop/met씮ile

RT-PCR SSCA, RE SSCA, RE

Ware et al., 1994 [25] Nafa et al., 1998 [43] Nafa et al., 1998 [43]

TM Luminal domain Luminal domain

181 182

Exon 6 Exon 6

1332 1348

G씮A AT씮C

444 450

trp씮stop Stop 455

Frameshift Nonsense Nonsense/ missense Nonsense Frameshift

SSCA, RE RT-PCR, SSCA

Nafa et al., 1998 [43] Bessler et al., 1994 [16]

Luminal domain B-LCL, luminal domain

183

Exon 6

1355

452

Stop 464

Frameshift

HA, SSCA

Nafa et al., 1998 [44]

184 185 186


1375 1381 1442

Ins AA-dup 32 nt (1324– 1355) Del G C씮T C씮T

459 481 481

Stop 491 arg씮trp ser씮phe

Frameshift Missense Missense

RT-PCR HA, SSCA Sequencing

Ostendorf et al., 1995 [37] Araten et al., 1999 [53] Nafa et al., 1998 [44]

6 6 6 6

Del ex 5

Healthy donor

39

40

Luzzatto and Nafa

TABLE 2-4 Summary of Molecular Abnormalities in the PIG-A Gene Type of mutation

Number reported

Obligatory loss of function (PIG-A⬚)

Large deletions Frameshift Deletion Insertion Del/insertion Ins/duplication Splice sitea In-frame deletionb Point mutations Nonsense Missense

3

3 99 (99/142 ⫽ 70%)

Total

65c,d 23e 7c 4 17c 4 23d,e 40c,e 186 f,g

17

23 — 142 (142/186 ⫽ 76%)

a

Alternative splicing producing a protein with partial activity cannot be excluded. b Some of these mutations can lead to a total loss of function. c Recurrent mutations: five deletions, one deletion/insertion, six splice sites mutations, four nonsense mutations, and four missense mutations (see Table 2-5). d Patient with non-Hodgkin lymphoma (NHL) treated with the anti-CD52 antibody. Campath-1H (two deletions, and two nonsense mutations). One patient had two PNH clones with different mutations [52]. e Mutations in seven healthy donors: one insertion, two nonsense mutations, and five missense mutations. One healthy donor has two different mutations [53]. f In total, 12 PIG-A mutations were found in 10 individuals without PNH. g The remaining 174 PIG-A mutations were identified in 146 patients with PNH.

Recurrent Mutations in the PIG-A Gene Sixteen recurrent mutations have been found in the PIG-A gene in patients with PNH. Four additional mutations were found to occur in patients with PNH and in individuals without PNH (Table 2-5, lines 31–32, 44–45, 85–88, and see below). All together, nine mutations have been found twice, one has been found three times, and three have been found four times.

PIG-A Mutations without PNH The monoclonal cytotoxic antibody anti-CD52 (Campath-1H) has been used in the treatment of patients with non-Hodgkin lymphoma [50] and with refractory chronic leukemia [51]. CD52 is a GPI-linked molecule, and it appears that the antibody is exerting selection in vivo for rare mutant cells with the PNH pheno-

41

2. Genetics of PNH

TABLE 2-5 Recurrent Mutations in the PIG-A Gene Substitution Number

Exon/intron

Nucleotide

Amino acid

Number of times

1 2 3 4 5 6 7 8 9 10 11 12 13

2 2 2 2 2 2 2 2 IVS4 IVS5 IVS5 6 6

163CAG씮TAG 229CGA씮TGA 294TAC씮TAA 298CAG씮TAG 336GC씮T 338T씮C 383CAT씮CGT 577delGTACT ⫹1g씮4 ⫹1g씮a ⫺2a씮g 1234delC 1309delC

55gln씮stop 77arg씮stop 98tyr씮stop 100gln씮stop Frameshift 113leu씮pro 128his씮arg Frameshift Splice Splice Splice Frameshift Frameshift

2 2a 2b 2 2 2 4 4c 4 3 2 2 2

a

One mutation was found in a PNH patient and one mutation was found in a healthy donor [30, 53]. b One mutation was found in a PNH patient and one mutation was found in a healthy donor [29, 53]. c Two mutations were found in patients with NHL treated with anti-CD52 antibody [52].

type, presumably arising from the lymphoma cell population [52]. Four PIG-A mutations (two frameshift and two nonsense mutations) have been identified in these patients (see Table 2-3, lines 87–88). It has been recently shown that cells that have the PNH phenotype often exist in normal individuals (at very low frequency, of the order of 20 per million). Four missense, one frameshift, and two nonsense mutations were found in seven cases out of 10 donors analyzed. Two of these mutations had been already reported in patients with PNH (Table 2-3, lines 31–32 and 44–45), confirming that PIG-A mutations are not sufficient for the development of PNH [53].

Structure–Function Relationships The increasing number of known missense mutations and in-frame deletions in the PIG-A gene may help to elucidate structure–function relationships in the PIG-A protein. Indeed, different single base pair substitutions at codon 183 of the PIG-A gene give rise to different PNH phenotypes (Table 2-3, lines 78–79) [42, 54]. A deletion of phenylalanine at position 151 was responsible of PNH type III phenotype [29]. In six PNH patients (four with complete deletion of GPI-anchored proteins, one with both types III and II, and nonspecified in one case), the mutations were found in the hydrophobic sequence (nt 1246–1326) that may act as a transmembrane domain [5]. Site-directed mutagenesis experiment suggests that codons H128, S129, and S155 are critical for the function of the PIG-A protein [55].

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TABLE 2-6 Clinical and Biological Features of Patients with PNH Hemolytic florid PNH

Hemolytic/ hypoplastic PNH

AA-PNH

Not specified

Total

I ⫹ III I ⫹ III ⫹ II I ⫹ II Not specified

25 12 1 15

— 1 — —

20 2 3 15

24 — 1 27

69 15 5 57

Total

53

1

40

52

146

Type of RBCs deficiency

Correlation between Phenotypes and Genotypes Among the 146 patients with PNH, 54 had ‘‘classic hemolytic’’ PNH, 40 had the ‘‘AA-PNH syndrome,’’ and in 52 cases the type was not defined (Table 2-6). Flow cytometry analysis of RBCs with anti-CD59 revealed only PNH type III cells in 69 cases, both PNH II and PNH III cells in 15 patients, and only PNH II cells in five patients. In the remaining 57 PNH patients the relevant information was not available. The majority of patients with PNH type III RBCs had a nonfunctional PIG-A protein [PIG-A⬚]. In each of 17 PNH patients, two different PNH clones with distinct mutations were identified [26, 29, 36, 41–43, 52]. In each of four PNH patients, more than two different PIG-A gene mutations were found [33, 34, 40, 44]. About a third of the patients who have more than one PNH clone (5/15 ⫽ 33%) have both PNH II and PNH III RBCs. Two PNH clones with distinct mutations were also found in individuals without PNH [52, 53].

Comparison of the Spectrum of Mutations in Three X-Linked Genes: PIG-A, FVIII, and G6PD Because PIG-A is an X-linked gene, it is of interest to compare the spectrum of its acquired mutations to that observed in inherited X-linked disorders. We have chosen deficiency of G6PD, the product of a gene that, like PIG-A is a housekeeping gene [56] and severe hemophilia A due to deficiency of factor VIII, the product of a tissue-specific gene [57]. Different patterns emerge from this comparative study (Fig. 2-12). In the case of factor VIII (leaving aside the inversion mutation now known to be responsible for more than 40% of all cases [58]), among the mutations causing severe hemophilia A (FVIII ⬍1%), there is a predominance of single base pair substitutions (61%) and a minority of frameshift mutations (18%). By contrast, in the PIG-A gene, the majority of mutations causing PNH are frameshift. Moreover, in the FVIII gene, about one-third of the single base substitutions causing severe hemophilia A were C씮T in a CpG dinucleotide; by contrast, there were no CpG affected by mutations in PIG-A. We do not have an explanation for this striking difference, but one possibility

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Figure 2-12 Mutation frequency in three X-linked disorders: PNH, G6PD deficiency, and hemophilia A. The spectrum of somatic mutations in the PIG-A gene in PNH is similar to the spectrum of inherited mutations in FVIII gene in severe hemophilia A; both produce no functional protein. By contrast, the mutations found in the G6PD gene in G6PD deficiency and in those found in the FVIII gene in moderate hemophilia A are virtually all missense mutations that will produce a protein with some residual activity. (Reprinted from [43].)

is less methylation in the PIG-A gene, at least on the active X chromosome. As for G6PD deficiency, practically all mutations are missense, and all of them have some residual enzyme activity; complete inactivation of the G6PD gene may not be compatible with human life [56]. The same might be true of PIG-A if the inactivating mutations were inherited, but the PIG-A mutations in PNH are instead acquired and confined to the hematopoietic system.

Conclusion In terms of the nosographic classification of human diseases, PNH is rather special and perhaps unique because, although it results from somatic mutations, it is not truly a malignant disease. As a consequence, the genetics of PNH is somewhat unconventional, relating not to inheritance but to somatic cell genetics. Indeed, formal genetic analysis has revealed in somatic cell hybrids that inactivating mutations of PIG-A are recessive, in keeping with the fact that this gene

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encodes one subunit of an enzyme. Biochemical genetics has identified a specific metabolic block in the GPI biosynthetic pathway, the transfer of GlcNAc onto phosphatidyl inositol. The molecular genetics of PIG-A has confirmed its sequence homology with a GlcNAc transferase and revealed in PNH patients a variety of mutations that cause either complete or partial loss of function of the gene product. Because these mutations are somatic, each mutation is rarely seen in more than one patient. Cytogenetic mapping to the X chromosome is highly significant because it easily explains how a single somatic mutation can produce the PNH phenotype. In view of the inactivation of the X chromosome in the somatic cells of females, X linkage of PIG-A does not entail a different prevalence of PNH in males vs. females, as it would in the case of an inherited condition. It is also pertinent to note that, if PNH were a malignant disorder, PIG-A would by definition be a tumor suppressor gene—the first such gene on the X chromosome. However, PNH is not a malignant disorder, and therefore there is no exception as yet to the rule that tumor suppressor genes are not sex linked. In the population genetics of somatic cells with PIG-A mutations, we can draw interesting parallels to the population genetics of organisms. We can imagine that PIG-A mutations continually arise in the bone marrow at a low frequency (as do germ-line mutations in a population). In most cases, the mutant clones remain very small and transient over time, because the mutation has taken place, not in a hematopoietic stem cell (HSC), but in a downstream progenitor cell whose progeny are quickly exhausted. If the mutation takes place in a true HSC, it may still be miniscule and temporary, because the loss of PIG-A function does not entail any growth advantage (just like neutral or deleterious germ-line mutations often have no future in a population). However, if the environmental circumstances change, and an autoreactive population of T cells or NK cells were to target a GPI or a GPI-linked molecule on HSCs, the PIG-A mutant clone would immediately find itself at a selective advantage, and hence would be favored for expansion (just as a mutant allele conferring resistance against an infectious agent would do in an animal population).

Acknowledgments We thank David J. Araten, Monica Bessler, Anastasios Karadimitris, and Rosario Notaro for many discussions and sharing of thoughts on PNH. We thank Marco Capasso for invaluable help in preparing Table 2-3.

References 1. Ham, T. (1937). Chronic haemolytic anaemia with paroxysmal nocturnal haemoglobinuria: Study of the mechanism of haemolysis in relation to acid–base equilibrium. N. Engl. J. Med. 217, 915–917. 2. Dacie, J. V., Israels, M. C. G., and Wilkinson, J. F. (1938). Paroxysmal nocturnal haemoglobinuria of the Marchiafava type. Lancet 1, 479–491. 3. Dacie, J. V. (1963). Paroxysmal nocturnal haemoglobinuria. Proc. R. Soc. Med. 56, 587–596. 4. Oni, B., Osunkoya, B. O., and Luzzatto, L. (1970). Paroxysmal nocturnal haemoglobinuria: Evidence of monoclonal origin of abnormal red cells. Blood 36, 145–152.

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23. Hillmen, P., Richards, S. J., Baker A. J., Rawston A. C., Crawford D. H., and Layton D. M. (1998). Congenital with paroxysmal nocturnal haemoglobinuria. Blood 92, 154a (Abstr. 619). 24. Lin, L. I., Liu, C. H., Chen, Y. C., Shen, M. C., Wang, C. H., Huang, Y. L., and Lin, J. K. (1997). PIG-A gene mutations in four Taiwanese patients with paroxysmal nocturnal haemoglobinuria following aplastic anaemia. Br. J. Haematol 97, 286–292. 25. Ware, R. E., Rosse, W. F., and Howard, T. A. (1994). Mutations within the PIG-A gene in patients with paroxysmal nocturnal hemoglobinuria. Blood 83, 2418–2422. 26. Yamada, N., Miyata, T., Maeda, K., Kitani, T., Takeda, J., and Kinoshita, T. (1995). Somatic mutations in the PIG-A gene found in Japanese patients with paroxysmal nocturnal hemoglobinuria. Blood 85, 885–892. 27. Pramoonjago, P., Wanachiwanawin, W., Chinprasertsak, S., Pattanapanayasat, K., Takeda, J., and Kinoshita, T. (1995). Somatic mutations of PIG-A in Thai patients with paroxysmal nocturnal hemoglobinuria. Blood 86, 1736–1739. 28. Pakdeesuwan, K., Siripanyaphinyo, U., Pramoonjago, P., Pattanapanyasat, K., Wilairat, P., Kinoshita, T., and Wanachiwanawin, W. (1997). Genotypic and phenotypic implications in paroxysmal nocturnal hemoglobinuria (PNH): A preliminary investigation. Southeast Asian J. Trop. Med. Public Health (Suppl) 3, 58–63. 29. Pramoonjago, P., Pakdeesuwan, K., Siripanyaphinyo, U., Chinprasertsuk, S., Kinoshita, T., and Wanachiwanawin, W. (1999). Genotypic, immunophenotypic and clinical features of Thai patients with paroxysmal nocturnal hemoglobinuria. Br. J. Haematol. 105, 497–504. 30. Savoia, A., Ianzano, L., Lunardi, C., De Sandre, G., Carotenuto, M., Musto, P., and Zelante, L. (1996). Identification of three novel mutations in the PIG-A gene in paroxysmal nocturnal haemoglobinuria (PNH) patients. Hum. Genet. 97, 45–48. 31. Nishimura, J. I., Inoue, N., Azenishi, Y., Hirota, T., Akaogi, T., Shibano, M., Kawagoe, K., Ueda, E., Machii, T., Takeda, J., Kinoshita, T., and Kitani, T. (1996). Analysis of PIG-A gene in a patient who developed reciprocal translocation of chromosome 12 and paroxysmal nocturnal hemoglobinuria during followup of aplastic anemia. Am. J. Hematol. 51, 229–233. 32. Du, T., Liu, E. K., Lu, Z. J., Shen, T., and Chang, C. N. (1997). Clinical features and PIG-A mutations of Chinese patients with paroxysmal nocturnal haemoglobinuria. Blood 90, 4b (Abstr. 2698). 33. Azenishi, Y., Ueda, E., Machii, T., Nishimura, J., Hirota, T., Shibano, M., Nakao, S., Kinoshita, T., Mizoguchi, H., and Kitani, T. (1999). CD59-deficient blood cells and PIG-A gene abnormalities in Japanese patients with aplastic anaemia. Br. J. Haematol. 104, 523–529. 34. Nishimura, J., Inoue, N., Wada, H., Ueda, E., Pramoonjago, P., Hirota, T., Machii, T., Kageyama, T., Kanamaru, A., Takeda, J., Kinoshita, T., and Kitani, T. (1997). A patient with paroxysmal nocturnal hemoglobinuria bearing four independent PIG-A mutant clones. Blood 89, 3470–3476. 35. Bessler, M., Mason, P. J., Hillmen, P., and Luzzatto, L. (1994). Mutations in the PIG-A gene causing partial deficiency of GPI-linked surface proteins (PNH II) in patients with paroxysmal nocturnal haemoglobinuria. Br. J. Haematol. 87, 863–866. 36. Bessler, M., Mason, P. J., Hillmen, P., and Luzzatto, L. (1994). Somatic mutations and cellular selection in paroxysmal nocturnal haemoglobinuria. Lancet 343, 951–953. 37. Ostendorf, T., Nischan, C., Schubert, J., Grussenmeyer, T., Scholz, C., Zielinska-Skowronek, M., and Schmidt, R. E. (1995). Heterogenous PIG-A mutations in different cell lineages in paroxysmal nocturnal hemoglobinuria. Blood 85, 1640–1646. 38. Miyata, T., Yamada, N., Iida, Y., Nishimura, J., Takeda, J., Kitani, T., and Kinoshita, T. (1994). Abnormalities of PIG-A transcripts in granulocytes from patients with paroxysmal nocturnal hemoglobinuria. N. Engl. J. Med. 330, 249–255. 39. Nagarajan, S., Brodsky, R. A., Young. N. S., and Medof, M. E. (1995). Genetic defects underlying paroxysmal nocturnal hemoglobinuria that arises out of aplastic anemia. Blood 86, 4656–4661. 40. Endo, M., Ware, R. E., Vreeke, T. M., Singh, S. P., Howard, T. A., Tomita, A., Holguin, M. H., and Parker, C. J. (1996). Molecular basis of the heterogeneity of expression of glycosyl phosphatidylinositol anchored proteins in the paroxysmal nocturnal hemoglobinuria. Blood 87, 2546–2557. 41. Endo, M., Beatty, P. G., Vreeke, T. M., Wittwer, C. T., Singh, S. P., and Parker, C. J. (1996). Syngeneic bone marrow transplantation without conditioning in a patient with paroxysmal nocturnal hemoglobinuria: In vivo evidence that the mutant stem cells have a survival advantage. Blood 88, 742–750.

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42. Nafa, K., Mason, P., Hillmen, P., Luzzatto, L., and Bessler, M. (1995). Mutations in the PIG-A gene causing paroxysmal nocturnal hemoglobinuria (PNH) are mainly of the frameshift type. Blood 86, 4650–4655. 43. Nafa, K., Bessler, M., Castro-Malaspina, H., and Luzzatto, L. (1998). Spectrum of somatic mutations in the PIG-A gene in paroxysmal nocturnal hemoglobinuria (PNH) includes large deletions and small duplications. Blood Cells Mol. Dis. 24, 370–384. 44. Nafa, K., Bessler, M., Deeg, H. J., and Luzzatto, L. (1998). New somatic mutation in the PIG-A gene emerges at relapse of paroxysmal nocturnal hemoglobinuria. Blood 92, 3422–3427. 45. Maugard, C., Margueritte, G., Tuffery, S., Rabesandratana, H., Demaille, J., and Claustres, M. (1997). Recurrent PIG-A mutation (IVS5⫹1G씮A) in a paediatric case of paroxysmal nocturnal haemoglobinuria: Detection by the protein truncation test. Br. J. Haematol. 98, 21–24. 46. Endo, M., Ware, R. E., Vreeke, T. M., Howard, T. A., and Parker, C. J. (1996). Identification and characterization of an inherited mutation of PIG-A in a patient with the paroxysmal nocturnal hemoglobinuria. Br. J. Haematol. 93, 590–593. 47. Rocchigiani, M., Lestingi, M., Luddi, A., Orlandini, M., Franco, B., Rossi, E., Ballabio, A., Zuffardi, O., and Oliviero, S. (1998). Human FIGF: Cloning, gene structure, and mapping to chromosome Xp22.1 between the PIGA and the GRPR genes. Genomics 47, 207–216 48. Pavlu, J., Mortazavi, Y., Tooze, J., Marsh, J. C., Gordon-Smith, E. C., and Rutherford, T. R. (1997). Paroxysmal nocturnal haemoglobinuria due to an 88 bp direct tandem repeat insertion in the PIG-A gene. Br. J. Heamatol. 98, 289–291. 49. Stafford, H. A., Nagarajan, S., Weinberg, J. B., and Medof, M. E. (1995). PIG-A, DAF and protooncogene expression in paroxysmal nocturnal hemoglobinuria-associated acute myelogenous leukaemia blasts. Br. J. Haematol. 89, 72–78. 50. Hertenstein, B., Wagner, B., Bunjes, D., Duncker, C., Raghavachar, A., Arnold, R., Heimpel, H., and Schrezenmeier, H. (1995). Emergence of CD52-, phosphatidylinositol-glycan-anchordeficient T lymphocytes after in vivo application of Campath-1H for refractory B-cell nonHodgkin lymphoma. Blood 86, 1487–1492. 51. Rawstron, A. C., Rollinson, S., Richards, S., Short, M. A., Morgan, G. J., Wright, D., Hale, G., and Hillmen, P. (1997). GPI-deficient cells (PNH phenotype) are present in most normal individuals. Blood 90, 273a (Abstr. 1202). 52. Merk, B., Hildebrand, A., Rojewski, M., Raghavachar, A., and Schrezenmeier, H. (1997). Heterogeneous PIG-A mutations in aplastic anemia, paroxysmal nocturnal haemoglobinuria and NonHodgkin’s lymphoma. Blood 90, 434a (Abstr. 1928). 53. Araten, D. J., Nafa, K., Pakdeesuwan, K., and Luzzatto, L. (1999). Clonal populations of hematopoietic cells with paroxysmal nocturnal hemoglobinuria genotype and phenotype are present in normal individuals. Proc. Natl. Acad. Sci. USA 96, 5209–5214. 54. Rollinson, S., Richards, S., Norfolk, D., Bibi, K., Morgan, G., and Hillmen, P. (1997). Both paroxysmal nocturnal hemoglobinuria (PNH) type II cells and PNH type III cells can arise from different point mutations involving the same codon of the PIG-A gene. Blood 89, 3069–3071. 55. Norris, E. R., Howard, T. A., Marcus, S. J., and Ware, R. E. (1997). Structural and functional analysis of the PIG-A protein that is mutated in paroxysmal nocturnal hemoglobinuria. Blood Cells Mol. Dis. 23, 350–360. 56. Beutler, E., Vulliamy, T., and Luzzatto, L. (1996). Hematologically important mutations: Glucose6-phosphate dehydrogenase. Blood Cells Mol. Dis. 22, 49–56. 57. Kemball-Cook, G., Tuddenham, E. G. D., and Wacey, A. I. (1998). The factor VIII structure and mutation resource site: HAMSTeRS version 4. Nucleic Acids Res. 26, 216–219. 58. Antonarakis, S. E., Rossiter, J. P., Young, M., Horst, J., de Moerloose, P., Sommer, S. S., Ketterling, R. P., Kazazian, H. H., Negrier, C., Vinciguerra, C., Gitschier, J., Goossens, M., Girodon, E., Ghanem, N., Plassa, F., Laqvergne, J. M., Vidaud, M., Costa, J. M., Laurian, Y., Lin, S. W., Lin, S. R., Shen, M. C., Lillicrap, D., Taylor, S. A. M., Windsor, S., Valleix, S. V., and Nafa, K. et al. (1995). Factor VIII gene inversion in severe hemophilia A: Results of an international consortium study. Blood 86, 2206–2212.


3 Hemolysis in PNH Charles J. Parker The Veterans Affairs Medical Center, Salt Lake City, Utah 84148 and Department of Medicine, Division of Hematology, University of Utah School of Medicine, Salt Lake City, Utah 84103

Introduction Hemoglobinuria is the defining clinical feature of paroxysmal nocturnal hemoglobinuria (PNH). Astute observations (by both patients and physicians) of the episodic, nocturnal characteristics of hemoglobinuria resulted in the recognition of the disease as an entity discrete from both paroxysmal cold hemoglobinuria and march hemoglobinuria, two similar syndromes also described in the nineteenth century. Investigations into the mechanism that underlies the hemoglobinuria account for most of the progress that has been made in defining the molecular basis of PNH. Ironically, the hemoglobinuria of PNH is an epiphenomenon that is a consequence of a more fundamental abnormality. Available evidence suggests that PNH arises in the setting of bone marrow injury because absence of one or more glycosylphosphatidylinositol (GPI)-anchored proteins is conditionally advantageous. In this case, the mutation that bestows the selective advantage (the exact nature of which is still enigmatic) affects a gene (PIG-A) that is essential for the synthesis of the membrane attachment moiety (the GPI anchor) of a functionally diverse group of proteins. As a result, these membrane proteins are deficient in PNH. Among the affected cellular constituents are two critically important complement-regulatory proteins [decay-accelerating factor, DAF (CD55) and membrane inhibitor of reactive lysis, MIRL (CD59)] that are expressed on all hematopoietic cells, including erythrocytes. Because the affected erythrocytes lack the membrane constituents (DAF and MIRL) that ordinarily prevent complement-mediated injury, they are abnormally susceptible to complement-mediated lysis when the alternative pathway of complement is activated, and the PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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resulting intravascular hemolysis gives rise to the characteristic hemoglobinuria of PNH. This chapter focuses on how the pathophysiological mechanisms underlying the hemolysis of PNH were uncovered and how studies directed to this end revealed the mechanisms by which normal human erythrocytes are protected against the lytic actions of complement.

Early History The early history of PNH has been recounted by Crosby [1] and is summarized in Chapter 1. These early studies [2–7] established a number of key points: red blood cells were dissolved within the vessels and not in either the kidney or in the urine; sleep played a critically important role in the hemolytic process; the condition was due to a defect in the red blood cells and it was thought that this defect made them unusually sensitive to the increase in blood acidity that occurred during sleep. Although the role of complement in the lytic process was suspected, interpretation of the experimental evidence failed to make it clear that a reaction with complement was responsible for lysis, and an accurate assessment of the process had to await the discovery of the alternative pathway of complement by Pillemer [7], some 40 years later.

The Acidified Serum Test of Ham In 1937, apparently unaware of previous studies by Strubing and by van den Berg, Thomas Hale Hamm [4] made the first of his extensive and seminal reports on PNH. Ham postulated that ‘‘Because of the elevation in the carbon-dioxide content of the arterial blood and the decrease in pH known to occur during sleep, it was suspected that a change in acid–base equilibrium was related to the increased hemoglobinemia of the patients during sleep.’’ Based on in vivo and in vitro studies, Ham [8, 9] made the following observations: (1) rapid hemolysis was observed when serum or plasma was acidified by using either equilibration with CO2 or addition of lactic acid; (2) the effects of CO2 were inhibited by addition of sodium bicarbonate; (3) hemolysis was observed if serum or plasma from normal volunteers was substituted for patients’ serum or plasma; (4) blood Group O red cells from a normal volunteer were not hemolyzed when resuspended in patients’ serum or plasma that was subsequently acidified with CO2 or lactic acid. These observations formed the basis for the acidified serum test, in which a patient’s washed red cells are incubated at 37⬚C in acidified normal serum or the patient’s acidified serum. The hemolysis characteristic of PNH is revealed by colorimetric analysis of the supernatent from the centrifuged sample. In 1939, Ham and Dingle [8] published a landmark paper that influenced the course of PNH research for the next 50 years. Subtitled ‘‘Certain Immunological Aspects of the Hemolytic Mechanisms with Special Reference to Serum Complement,’’ this publication suggested a novel mechanism by which the abnormal erythrocytes of PNH were lysed by an immune mechanism that was independent of antibody. Building on previous observations that the hemolysis of PNH cells

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was inhibited by heat-inactivation of serum or by the addition of certain salts that were known to inhibit complement [4, 5], they had systematically investigated each facet of the humoral immune system (antigen, antibody, and complement) that was known at the time. That no evidence of antibody was found either in patient serum or associated with patient red cells implied the existence of a novel pathway of immune lysis. They also demonstrated that PNH red cells were more susceptible to lysis than normal red cells when incubated with antibody (either rabbit anti-human RBC antiserum or isohemolysins) and human serum. Lysis of PNH and normal red cells did not differ, however, in nonimmunological systems (saponin, sodium taurocholate, and hypotonic sodium chloride). The authors demonstrated that the characteristics of lysis of PNH red cells in acidified serum were markedly different from those of lysis of normal cells using isohemolysins (i.e., anti-blood group A, and anti-blood group B antibodies), again, consistent with a novel mechanism of hemolysis different from known antibody-initiated processes. Their finding that antibody did not participate in the lytic process was particularly remarkable because such a process was unprecedented. The authors were equally rigorous in their investigation of the role of complement in acidified serum lysis. They used available methods to increase or decrease complement concentration, inhibit complement, and remove or inactivate fractions or components of complement [8]. Their observation that PNH erythrocytes were more susceptible to antibody-initiated lysis when human serum was used as the source of the lytic substance was subsequently exploited by others to characterize the phenotypic mosaicism of the erythrocytes that is a defining feature of PNH [10, 11]. The final conclusion of Ham and Dingle that ‘‘The serum factor essential for hemolysis was closely associated with, if not indistinguishable from complement or alexin of human serum’’ suggests that the authors believed that the lytic substance of serum was indeed complement.

Relationship of Hemoglobinemia and Hemoglobinuria to Sleep Before leaving Ham’s work it seems appropriate to make a final comment on the relationship between sleep and hemoglobinuria, as it was this clinical feature that fascinated early investigators and distinguished PNH from other hemolytic anemias. The symptom is also important to patients because it is a disconcerting reminder of the chronic nature of their illness and often serves as a dreaded harbinger of an exacerbation of the disease. Finally, general audiences remain enamored with this symptom, and following any lecture on PNH, a question that invariably arises is ‘‘Why does the hemolysis occur at night?’’ Ham hypothesized that the hypoventilation associated with sleep is the critical factor that is responsible for the associated increase in hemoglobinemia. He first demonstrated that hemoglobinuria markedly increased during sleep. The daily excretion of hemoglobin was quantitated and divided into periods of waking and sleep. The analysis was performed for 10 consecutive days, and the results clearly showed a marked increase in hemoglobinuria during sleep. Next, Ham investigated the relationship between sleep and hemoglobinemia [9]. When the patient was kept awake for 27 consecutive hours, the normal pattern of increased nocturnal hemoglobinemia was no longer observed. Even if he then altered the

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pattern of sleep so that the patient slept during the day, hemoglobinemia was again observed to increase; bed rest without sleep did not result in increased hemoglobinemia. Thus, paroxysmal nocturnal hemoglobinuria is something of a misnomer—increased hemolysis is related to sleep, rather than to time of day. To examine the effects of respiration on hemoglobinemia, Ham used a Drinker respirator to artificially control both the respiratory rate and volume [9]. Hyperventilation during 6 h of sleep produced alkalosis and was accompanied by no increase in either hemoglobinemia or hemoglobinuria (Table 3-1); during sleep without hyperventilation, a more acid pH was observed, accompanied by an increase in both hemoglobinemia and hemoglobinuria. Results at variance with those of Ham were reported by Crosby [12] in 1953. Crosby concluded that ‘‘The ‘acid shift’ undoubtedly contributes to the ‘nocturnal’ nature of the disease, but it is not the only pathological mechanism’’ [12]. Others have also questioned whether acidification of plasma is an important factor in the relationship between sleep and hemoglobinemia and not all patients with PNH are observed to have an increase in hemoglobinemia during sleep [13]. Conceivably, among patients who have a nocturnal increase in hemoglobinemia, the blood may be acidic in localized areas where the rate of flow is slowed during sleep, but the pH of arterial blood sampled from an extremity may not reflect such heterogeneity. In support of this hypothesis, Stru¨bing [3] noted that heavy exercise or overwork produced an attack of hemoglobinuria in his patient the subsequent night. He thought that accumulation of lactic acid from the previous days’ exertion contributed to the acid environment needed to induce hemolysis. Eighty-five years later, Blum and colleagues [14] reported that the exacerbation of hemolysis noted after strenuous exercise is associated with an increase in plasma lactic acid concentration and with a concordant reduction in the pH of the blood. The basis of the increase in hemoglobinemia and hemoglobinuria that occurs during sleep is not an area of active investigation. Based on available evidence, it is impossible to conclude unequivocally that the nocturnal aspect of the hemoglobinuria is due to acidification of the plasma that results from accumulation of CO2 as a consequence of hypoventilation associated with sleep. The question of the relationship between hemolysis and sleep is interesting from a historical perspective, and, more importantly, a better understanding of the relationship

TABLE 3-1 Effect of Alkalosisa on Hemoglobinemia

Condition

Plasma hemoglobin, (mg/100 cc)

Urine hemoglobin

pCO2

pH

O2 saturation (%)

Awake and ambulatory Natural sleep; respiration normal Natural sleep; hyperventilation

95–150 230 140–150

⫹⫹ to ⫹⫹⫹ ⫹⫹⫹⫹ ⫹⫹⫹

42.0 28.0

7.3 7.47

93.9 97.95

a

Alkalosis was produced by hyperventilation during natural sleep.

53

3. Hemolysis in PNH

between sleep and hemolysis might suggest more specific therapy for this troubling symptom.

Discovery of the Alternative Pathway of Complement The decade that followed the 1939 publication of Ham and Dingle [8] produced only limited progress toward defining the basis of the abnormal susceptibility of PNH erythrocytes to hemolysis in acidified serum. The role of complement and even the role of divalent cations in the hemolytic process associated with PNH was debated into the mid-1950s [15]. In 1951, Harris and colleagues [16] observed that the process responsible for the lysis of PNH erythrocytes in vitro differed from hemolytic complement by its dependence on magnesium and its inhibition by calcium. At the time, the findings of Harris et al. appeared to represent only a minor incremental advance; in retrospect, however, the observation that the hemolysis of PNH was a magnesium-dependent phenomenon can be seen as a pivotal. In 1954, Pillemer, (a co-author with Hevrd) [17] reported the serendipitous isolation of properdin during an attempt to isolate one of the components of C’3 (C’3 of complement actually consisted of several discrete proteins). A year earlier [18a, 18b], Pillemer had noted the striking resemblance between the mechanism of lysis of PNH erythrocytes and the inactivation of complement by zymosan: both reactions required magnesium and serum components resembling complement, and both proceeded optimally at pH 7, but neither system required a specific antibody. Subsequent experiments showed that the properdin system was involved in the lysis of PNH erythrocytes in acidified serum [17]. Moreover, Hinz, Jordan, and Pillemer [19] showed that serum depleted of properdin (RP) had no hemolytic activity for PNH erythrocytes in acidified serum, whereas RP was fully active in mediating lysis of red cells in antibody-dependent systems; repleting the RP with isolated properdin restored its capacity to support lysis of PNH red cells in acidified serum (Table 3-2). Initially, properdin was envisioned as functioning in a manner analogous to antibody (i.e., its binding to the cell surface was thought to initiate activation of complement). However, in contrast to antibody (a component of acquired immunity), properdin was assigned by Pillemer as the central element of the

TABLE 3-2 The Relation of Serum Properdin Level to Hemolysis of PNH Erythrocytes

Serum

Properdin content (units/ml)

Hemolysis of PNH erythrocytes (%)

Normal—No. 1 Normal—No. 2 RP RP ⫹ 3 units properdin RP ⫹ 5 units properdin

4–8 1–2 0 3 5

33 14 0 23 36

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innate or natural immune system [17]. This recognition of a mechanism for complement activation that did not require antigenic stimulation was a brilliant conceptual leap. According to Pillemer’s hypothesis, properdin is present in the serum constitutively (i.e., synthesis of the protein does not depend on an antigenic stimulus), and binding is determined, not by recognition of a specific antigen, but by the innate biochemical properties of the cell surface. Thus, constituents of self (e.g., erythrocytes) remain unharmed because properdin has no natural affinity for host cells. In contrast, the system protects against infections because properdin binds to invading organisms through a system that recognizes the foreign composition of the cell membrane surface. Although the details of the mechanism of activation of the alternative pathway are now known to differ from this model (discrimination of self and nonself is mainly determined by the factor H component of the alternative pathway), the basic concept suggested by Pillemer was correct. In an insightful review of PNH published in 1963, Dacie [20] stated that the conclusion that the properdin system was responsible for the lysis of PNH erythrocytes in acidified serum ‘‘seems generally to have been widely accepted.’’ He then proposed that the abnormal sensitivity of PNH erythrocytes was due to abnormality in the surface of PNH red cells. Dacie speculated ‘‘it is possible that this leads to the adsorption of the properdin complex, while normal cells by virtue of their normal surface fail to do this to a significant degree. The fixation of the complex to the cell surface then leads enzymatically to haemolysis. Presumably it is the complement complex which actually brings about the lysis, with properdin, by fixing the complement to the cell, playing a role analogous to that played by classical amboceptor.’’ The wide acceptance of the role of the properdin system or alternative pathway in the lysis of PNH erythrocytes was the result of two factors. First, the data from the studies of Pillemer, Hinz, and colleagues [17, 19] were compelling. Second, the hypothesis fit precisely with what was known about acidified serum lysis of PNH erythrocytes (magnesium along with components resembling complement were required, but antibody was not involved). Many within the immunological community doubted the existence of the properdin pathway, however, due to the fact that preparations of purified properdin were found to contain small amounts of natural antibodies to yeast and bacteria (protein purification was in its infancy at the time and isolation to homogeneity was in most cases technically impossible). Pillemer’s work was cut short by his death, in 1957, of a barbiturate overdose that was ruled a suicide. (He was apparently depressed over the failure of other complement investigators at a meeting to accept his properdin system.) Ten years later, the alternative pathway was rediscovered by others, and by 1980 all of the components of the system had been identified and isolated. To appreciate the basis of the hemolysis of PNH, it is necessary to understand the mechanism of action of the alternative pathway. A comparison of the alternative pathway with the classical pathway is required because some important features of PNH were elucidated by careful analysis of the susceptibility of PNH erythrocytes to complement-mediated lysis initiated by antibody. Further, a clinical laboratory test (the sucrose lysis test of Hartmann [21, 22]) that is still

3. Hemolysis in PNH

55

Figure 3-1 Schematic representation of the classical and alternative pathways of complement. The stability (—) of the C1 complex (C1q, C1r, and C1s) is calcium (Ca2⫹) dependent. Binding of C2b to C4b in the classical pathway and binding of factor B to C3b in the alternative pathway are magnesium (Mg2⫹) dependent (⫹). The * symbol indicates enzymatic activity. The erythrocyte membrane proteins that regulate complement are shown below the complex that they regulate, and the membrane regulatory factors that are deficient on PNH erythrocytes are denoted by bold type. The serum factors that regulate the classical pathway, the alternative pathway, and the membrane attack complex (MAC) are shown above the complexes that they regulate.

used in the diagnosis of PNH depends, at least in part, on activation of the classical pathway [23].

The Classical Pathway of Complement1 The classical pathway (Fig. 3-1) is activated on erythrocytes when the first component of complement (C1) binds to the Fc portion of IgG or IgM. Antibodies of the IgA, IgD, and IgE classes lack the binding site for C1 and thus are unable to activate complement. Of the IgG subclasses, IgG3 binds C1 most efficiently, followed in order of efficiency by IgG1 and IgG2. IgG4 interacts poorly with C1 and appears to lack the capacity to activate the classical pathway [24]. 1

The description of the complement system was modified from Parker, C. J., and Foerster, J. (1998). Mechanisms of immune destruction of erythrocytes. In ‘‘Wintrobe’s Clinical Hematology’’ (G. R. Lee, J. Foerster, J. Lukens, F. Paraskevas, J. P. Greer, and G. M. Rodgers, Eds.), 10th Ed., pp. 1192–1197. Williams & Wilkins, Philadelphia, with copyright permission of Williams & Wilkins.

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C1 is actually composed of three subunits (C1q, C1r, and C1s). The stoichiometry is as follows: C1q, C1r2, C1s2. The C1q component binds weakly to the C1r2C1s2 tetramer, and the structural and functional integrity of the C1 complex is calcium-dependent [25] (Fig. 3-1). The binding constant for this reaction (5 ⫻ 107 M⫺1) is such that approximately 80% of the C1q and C1r2C1s2 subunits is combined as C1 molecules in the plasma [26, 27]. C1q is the subunit of the complex that binds to the Fc portion of IgG or IgM. The ultrastructure of C1q is complex and distinctive [24, 28, 29]. The molecule is composed of six globular heads extending from a collagen-like stem with the six stems joined together in a bunch (under electron microscopy, the side view of the molecule looks remarkably similar to a bouquet of tulips). Each of the six globular heads of C1q has a binding site that is specific for a region in the Fc portion of the immunoglobulin molecule [24]. Binding of a single head, however, is weak and lasts only a fraction of a second. Firm binding lasting several minutes requires the interaction of at least two of the heads. While each IgG molecule has two C1q binding sites (one in each of its two Fc domains), C1q cannot bind firmly to a single IgG molecule because the binding sites in IgG are located on opposite sides of the Fc moiety. Inasmuch as the C1q molecule is relatively inflexible, two of the globular heads cannot engage both of the Fc binding sites simultaneously. For IgG to activate the classical pathway, at least two molecules must be bound in close proximity (within 30–40 nm) [28, 30, 31]. This requirement means that the number of copies of antigen per cell is an important determinant of complement activation. In some cases, however, antigens that are present in a relatively low number of copies may be able to support complement activation, if they are free to move within the plane of the membrane so that patches or clusters can form. The relatively low density of the antigen, its even distribution over the entire membrane surface, and its inability to patch due to constraints imposed by its structure and interactions with other membrane constituents probably account for the lack of complement activation induced by anti-Rh antibodies [32, 33]. Thus, as noted by Dacie in 1949 [10], PNH erythrocytes are no more susceptible to lysis initiated by anti-Rh antibodies than are normal erythrocytes (in fact, anti-Rh antibodies do not induce hemolysis of either normal or PNH erythrocytes). The efficiency with which IgG is able to activate the classical pathway is also enhanced when an immune reaction generates polyclonal antibodies that recognize different epitopes on the same antigen. Under these conditions, the stringent spacial requirements for C1q binding to Fc may be met when two or more molecules of IgG bind to different epitopes on a single antigenic structure. In contrast to IgG, IgM is a highly efficient activator of the classical pathway. The pentameric structure of IgM means that five Fc moieties are present in the same molecule with at least three (and probably all five) C1q binding sites available [34]. Thus, a single molecule of IgM has the capacity to activate the classical pathway. However, the C1q binding sites are not exposed unless the IgM molecule assumes the ‘‘staple’’ form. In order to assume this configuration, at least two of the subunits of the IgM molecule must engage the cellular antigen [35]. Thus, antigen density also impacts on the capacity of IgM to activate comple-

3. Hemolysis in PNH

57

ment. When free in plasma, IgM does not activate complement because it exists in a star-shaped, planar form. Under these conditions, the C1q binding sites are not exposed. Interaction with antigen causes the F(ab⬘)2 arms to bend so that they are at right angles to the central Fc core of the molecule (the staple configuration). This conformational change exposes the C1q binding sites, allowing complement activation to proceed. Both C1r and C1s are serine proteases that exist in their native state as zymogens (inactive proenzymes). As noted above, the integrity of the C1r2C1s2 complex is dependent on calcium [36] (Fig. 3-1), and the C1s molecules are located at either end of the linear tetramer with the two C1r molecules in the middle [24]. The primary structure of the two proteins is very similar, suggesting that they arose by gene duplication. Enzymatic activity is expressed following proteolytic cleavage of the zymogens. Apparently, binding of C1q to immunoglobulin induces a conformational change in C1 that causes autoactivation of the two C1r molecules [36]. The precise mechanism by which autoactivation occurs remains obscure, but available evidence suggests that as each C1r molecule undergoes autoactivation, it cleaves its partner within the tetramer. By enzymatic cleavage, the two proteolytically activated C1r molecules subsequently activate their contiguous C1s partners [24]. Binding of C1 is rapid, but activation is a relatively slow process [36], probably because not all C1 binding sites support activation of C1. Binding of C1 to immunoglobulin is a crucial amplification step in the generation of the classical pathway cascade because activated C1s has the capacity to proteolytically cleave many molecules of C4 and C2. To counter this enormous destructive capacity, elaborate safeguards exist to restrict the activity of complement. These check points usually involve inhibition of the formation or activity of an enzymatic step in the pathway so that amplification is dampened. The first such complement regulatory factor is C1 inhibitor (Fig. 3-1). This heavily glycosylated plasma protein binds in a 1 : 1 stoichiometric relationship with activated C1r and C1s, forming an extremely stable (probably covalent) complex. Binding of C1 inhibitor causes activated C1r and C1s to dissociate from the C1 complex, leaving C1q bound to immunoglobulin [24]. Because the rate of reaction of C1 inhibitor with activated C1r and C1s is so rapid, the half-life of activated C1 in the plasma in only 10–20 s [37]. C1 inhibitor is a member of the superfamily of serine protease inhibitors (serpins), and it also has the capacity to block the activities of plasmin, kallikrein, factor XIa, and factor XIIa [32]. (Deficiency of C1 inhibitor underlies the clinical disorder angioedema, a disease that exists in both inherited and acquired forms [38].) The natural substrates for activated C1 are C4 and C2. Activated C1s cleaves C4 near the amino terminus of the 움 chain of this disulphide-linked heterotrimer [39]. The smaller cleavage fragment, designated C4a, is a weak anaphylatoxin [40]. The larger fragment, C4b, contains an internal thioester bond that is exposed when C4 is cleaved by C1s [41]. By acyl transfer, this thioester group located in the 움 chain can form either an ester or an imidoester bond with the immunoglobulin molecule or with an erythrocyte membrane glycoprotein (in particular, glycophorin A) [42, 43]. The half-life of activated C4 is extremely short (앑60 애s by analogy with nascent C3b, see below). During this period, the activated protein can

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Charles J. Parker

diffuse about 40 nm. Thus, C4b is found in clusters around the immunoglobulin–C1 complex. The covalent attachment of C4b serves as the nidus for the formation of the C3 convertase of the classical pathway by providing a magnesium-dependent binding site for C2, a zymogen belonging to the family of serine proteases (Fig. 3-1) [44]. C2 that is bound to C4b at a distance no greater than 60 nm from an immunoglobulin-activated C1 complex is cleaved by C1s into fragments of C2a and C2b. The C2b fragment remains bound and functions as the enzymatic subunit of the classical pathway C3 convertase (there is no universal agreement on the nomenclature of the C2 fragments, and many investigators continue to use the original nomenclature in which the enzymatic subunit of C2 was designated C2a). No definite biological function has been attributed to nonenzymatic C2a peptide. The activity of the C3 convertase of the classical pathway is regulated by its intrinsic instability and by both plasma and membrane proteins (Fig. 3-1). The plasma constituent. C4b-binding protein, restricts convertase activity by either binding to C4b and inhibiting the subsequent binding of C2 or by accelerating the decay of formed C4b2b complexes, or both [45–47]. C4b-binding protein also serves as a cofactor for the enzymatic degradation of C4b by the plasma serine protease, factor I [47–49]. C4b-binding protein has an interesting structure, in that it is composed of seven identical chains that are linked by disulfide bonds [50]. Electron micrographs show a spider-like structure consisting of seven elongated subunits (resembling tentacles) linked to a small central body [51]. Approximately 50% of C4b-binding protein in plasma is complexed with protein S, the vitamin K-dependent protein C cofactor that plays an important role in hemostasis [50]. The activity of the classical pathway C3 convertase (C4b2b) is also regulated by two erythrocyte membrane proteins, complement receptor type 1 (CR1, CD35) and decay-accelerating factor (DAF, CD55) (Figs. 3-1 and 3-2). CR1 regulates the convertase activity in much the same way as C4b-binding protein (i.e., it binds to C4b preventing formation of the complex, accelerates the decay of the C4b2b complex, and serves as a factor I cofactor) [52–54]. In contrast, DAF lacks factor I cofactor activity, and it does not appear to block convertase formation. Rather (as its name suggests), DAF restricts convertase activity by accelerating the decay of the C4b2b complex [55–57]. Membrane cofactor protein (MCP, CD46) is another cellular protein that inhibits the complement activation. Like CR1, MCP (CD46) serves as a cofactor for the factor I-mediated cleavage of C4b (and C3b) [57, 58]. MCP (CD46), however, is not an inhibitor of immune hemolysis because it is not expressed on erythrocytes. CR1 and DAF share a common structural motif consisting of cysteine-rich units of approximately 60 amino acids, called short consensus repeats (SCR). DAF contains four SCRs while CR1 contains 30 such units (this number actually varies depending upon the isoform). This structural feature (the SCR) is also shared by other complement proteins [factor H, C4b-binding protein, MCP (CD46), and complement receptor type 2 (CR2, CD21)]. All of these proteins interact with C3b or C4b or both. The genes for these proteins have been localized to the long arm of chromosome 1 (band q32), and together, this gene family is referred to as the regulators of complement activation (RCA) [59]. CR1 is a

3. Hemolysis in PNH

59

Figure 3-2 Erythrocyte complement-regulatory proteins. The ectoplasmic portions of CR1 (CD35) and DAF (CD55) are comprised in large part of a series of SCRs ( ) consisting of 앑 60 amino acids each. The ectoplasmic portion of MIRL (CD55) ( ) is structually unrelated to CR1 and DAF. CR1 is an integral membrane protein with transmembrane ( ) and cytoplasmic ( ) domains in addition to an ectoplasmic domain. In contrast, both DAF and MIRL are anchored by a glycosyl phosphatidylinositol moiety, consisting of phosphatidylinositol ), glucosamine ( ), and three mannose molecules ( ). This unit is linked to the COOH terminus of the ecto( plasmic domain by ethanolamine, and the diacyl portion of phosphatidylinositol ( ) anchors the molecule into the lipid bylayer. [Reproduced from Parker, C. J., and Foerster J. (1998). Mechanisms of immune destruction of erythrocytes. In ‘‘Wintrobe’s Clinical Hematology’’ (G. R. Lee, J. Foerster, J. Lukens, F. Paraskevas, J. P. Greer, and G. M. Rodgers, Eds.), 10th Ed. pp, 1195. Williams & Wilkins, Philadelphia, by copyright permission of Williams & Wilkins.]

type I integral membrane protein while DAF is anchored to the cell through a glycosylphosphatidylinositol (GPI) moiety (Fig. 3-2). Like all GPI-anchored proteins, DAF is deficient on the erythrocytes from patients with PNH, and the deficiency of DAF accounts in part for the complement-mediated lysis that is the clinical hallmark of the disease. DAF (CD55) is discussed in more detail below. The classical pathway C5 convertase is formed when C3 in the fluid phase is cleaved into C3a and C3b fragments by the C2b component of the C4b2b complex (Fig. 3-1). The 77 amino acid C3a molecule that is liberated from the amino-terminal end of the 움 chain of C3 as a consequence of cleavage by C2b is an anaphylatoxin that is 앑10-fold more potent than C4a, but 앑200-fold less potent than C5a [40, 60]. The receptor for C3a has been cloned [61, 62]. Although the C3a receptor is structurally and functionally similar to the C5a receptor, the tissue distribution of the two anaphylatoixin receptors appears to be different. Whether the C3a receptor is also the receptor for C4a is under investigation. Like C4b, C3b contains an internal thioester bond that mediates the covalent attachment of the molecule to immunoglobulin or cell surface constituents [63– 65]. Thus, the C3b that becomes membrane bound is found in clusters around the C4b2b complex, because the extremely short half-life of the binding site (앑60 애s) limits the distance that nascent C3b can diffuse before it becomes inactive [66]. While only 10–15% of the C3b that is generated by C4b2b becomes cell bound, this step is nonetheless an important part of the amplification process because each convertase mediates deposition of 앑200 molecules of C3b [67]. The C5 convertase is regulated by the same plasma and cellular proteins that control the activity of the C3 convertase (Fig. 3-1). In the case of the C5

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convertase, CR1 also binds to C3b and serves as a cofactor for factor I mediated degradation of C3b to iC3b and subsequently to C3dg [68, 69]. This latter C3 degradation fragment is the predominant form of C3 found on circulating erythrocytes in patients with immune hemolytic anemias involving complement activation (e.g., chronic cold agglutinin disease) [70]. In addition to CR1, the plasma protein factor H serves as a cofactor for factor I-mediated degradation of C3b to iC3b. Under physiological conditions, however, factor H does not support the factor I mediated cleavage of iC3b to C3c and C3dg [68]. Factor H, a single chain glycoprotein consisting of 20 SCR units [71], is an important regulator of the C3 and C5 convertases of the alternative pathway (discussed below). C2b is also the catalytic subunit of the classical pathway C5 convertase (C4b2b3b, Fig. 3-1) [72]. In this case, cell-bound C3b serves as the attachment site for C5, thereby positioning it for enzymatic activation by C2b. Nascent C5b is generated when C2b cleaves a 74 amino acid peptide (C5a) from the amino terminal portion of the 움 chain of C5 [73, 74]. In addition to being a potent anaphylatoxin, C5a is an important component of the immune process that mediates a variety of proinflammatory events through binding with a specific G protein coupled receptor expressed on a variety of tissues [75]. Unlike C4b and C3b, C5b does not contain an internal thioester bond, and thus it does not bind covalently to the cell membrane [41]. Rather, C5b serves as the nidus for the formation of the membrane attack complex (MAC). The number of C5b molecules that are activated by the classical pathway C3 convertase is greatly constrained: only C5 that is bound to a C3b molecule that is in close proximity to a C4b2b complex can be cleaved. Thus only two or three molecules of C5b may be generated by each C4b2b complex. The functional half-life of C5b is about 2 min, and it appears to undergo irreversible inactivation while still bound to C3b [72].

The Membrane Attack Complex (MAC) After the generation of C5b, no further enzymatic activity is involved in either the formation or the cytolytic activity of the MAC that consists of C5b, C6, C7, C8, and C9 (C5b-9). Components C6, C7, and C9 are single-chain polypeptides [76–82]. As discussed above, C5b is a disulfide-linked heterodimer that is the product of a single gene. C8 consists of three nonidentical peptide chains [83]. The 움 and 웂 chains comprise a disulfide bonded unit that is noncovalently associated with the 웁 chain. Apparently the three polypeptide chains that comprise C8 are the products of separate genes [84]. There are structural and antigenic similarities among four of the components (C6, C7, C8, and C9), and at the nucleotide level, modest homology among C7, C8움, C8웁, and C9 [66]. Formation of the MAC is initiated when nascent C5b (that is still complexed with C3b) binds specifically to C6, forming a stable, hydrophilic complex [85, 86]. Binding to the C5b6 complex induces a conformational change in C7 to expose a labile membrane-binding site and cause the molecule to undergo a hydrophilic–amphiphilic transition [87, 88]. This process allows the trimolecular C5b67 complex to integrate into the lipid bilayer of the erythrocyte. Once inserted into the cell, the complex is stable. Binding of C8 to the C5b67 complex is mediated through the 웁 chain that has a specific recognition site for C5b [89].

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61

Binding to C5b67 induces a conformational rearrangement that allows the 움 chain of C8 to insert into the hydrophobic core of the lipid bilayer and exposes a single binding site for C9. Binding to C8움 within the C5b–8 complex [90] causes C9 to unfold. This conformational change exposes hydrophobic regions that allow the molecule to insert into membrane and exposes a binding site for an additional C9 molecule. Binding to C5b–91 causes the second C9 molecule to undergo the same conformational change as the first C9 molecule, and in this way multiple molecules of C9 become incorporated into the MAC (C5b–9n) [66]. The C9 molecules undergo polymerization to form a ring-like structure that appears as a classical doughnut lesion on electron microscopy [91, 92]. Individual MACs are heterogeneous in size due to differences in C9 composition [93–95]: the stoichiometry of the MAC is C5b1, C61, C71, C81, C9n, where n ranges from 2 to 18, depending upon experimental conditions [66, 93, 94]. On normal human erythrocytes that have undergone hemolysis, the average C8:C9 ratio is 1:3, whereas on PNH erythrocytes that have undergone lysis, the average C8:C9 ratio is 1:6 [96]. These results predicted that PNH erythrocytes were deficient in a membrane constituent that regulates the formation of the MAC (see below). Disruption of the integrity of the lipid bilayer of the erythrocyte by the MAC creates an osmotic gradient because small ions can traverse the damaged membrane but large cytoplasmic components such as hemoglobin cannot. The consequent inflow of water causes the cell to expand rapidly, and if damage is great enough, the cell ruptures and releases hemoglobin into the plasma (a process called colloid osmotic lysis). The mechanism by which the MAC produces cell lysis is debated. According to the ‘‘leaky patch’’ hypothesis, insertion of the hydrophobic elements of the MAC causes local disruption of the integrity of the phospholipid bilayer [97]. Proponents of the competing ‘‘pore’’ hypothesis argue that the polar surfaces of the MAC components aggregate, forming a hydrophilic channel through the membrane [98]. Both processes may be operative, depending upon the experimental conditions. A number of plasma proteins, including S protein (vitronectin) [99], and apolipoproteins (particularly clusterin), have been shown to inhibit activity of the MAC in vitro (Fig. 3-1) [100–104]. The importance of these plasma constituents in regulating the lytic actions of complement in vivo, however, has not been unequivocally established. Erythrocytes are protected from lytic action of the MAC by membrane inhibitor of reactive lysis (MIRL, CD59) [105, 106]. This GPI-anchored protein (Fig. 3-2) inhibits MAC formation primarily by binding to C8, thereby restricting the subsequent binding and polymerization of C9 [107–109]. The importance of MIRL in vivo was illustrated by a patient with an isolated deficiency of this complement regulatory protein [110], who had a syndrome characterized by recurrent episodes of intravascular hemolysis clinically indistinguishable from PNH. MIRL (CD59) is discussed in greater detail below.

The Alternative Pathway of Complement Activation of the classical pathway is highly specific in that antibody is required to initiate the process. The system is quiescent unless erythrocyte antigens become

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targets for immune attack as a consequence of some pathological process— autoimmune, alloimmune, or isoimmune. Therefore, under normal physiological conditions, host erythrocytes do not require protection against the classical pathway because antibodies against self do not arise. In contrast to the classical pathway, there is no initiating factor for the alternative pathway—the alternative pathway is in a state of continuous, low-grade activation. Thus, the burden of avoiding attack by the alternative pathway is borne directly by host cells that rely on membrane factors to restrict the activity of the system. Because they are always exposed to complement activation by the alternative pathway, erythrocytes have evolved specific cell surface constituents (DAF and MIRL) that protect them against complement-mediated destruction. That normal erythrocytes never undergo immune destruction mediated by the alternative pathway is a testament to the effectiveness of these two proteins. On the other hand, the chronic complement-mediated intravascular hemolysis of PNH is mediated by the alternative pathway, underscoring the essential role of DAF and MIRL in protecting normal erythrocytes from injury. As antibody is not required for activation, the alternative pathway functions as the first line of defense against infections in the nonimmune host [111] (the concept of natural or innate immunity as envisioned by Pillemer [17]). The alternative pathway is activated by certain microorganisms because the biochemical properties (e.g., the sialic acid content) of these potential pathogens promote the formation and stability of C3 convertase. On alternative pathway activators, the binding of the catalytic subunit (factor B) of the C3 convertase is favored over the binding of the convertase inhibitory protein (factor H). Erythrocytes are exposed to the same plasma factors as invading organisms, and yet normal red cells are completely resistant to injury mediated by the alternative pathway. Erythrocytes serve as a paradigm for understanding the elaborate mechanisms that have evolved to protect host cells against the untoward consequences of alternative pathway activation. In plasma, activation of the alternative pathway is initiated by direct formation of the C3 convertase, consisting of hydrolyzed C3 and activated factor B [112]. The activity of the system is kept at a low level because, in the fluid phase, the formation and stability of the amplification C3 convertase are tightly controlled by the endogenous plasma regulatory protein, factor H (Fig. 3-1). Nonetheless, some low-grade activation does occur because the internal thioester in native C3 is subject to spontaneous hydrolysis. Nascent C3 ⭈ H2O undergoes a conformational change that transiently exposes a magnesium-dependent binding site for factor B [112]. The catalytic subunit of the alternative pathway C3 convertase (Bb) is generated when factor B that is bound to C3 ⭈ H2O is cleaved by factor D into Ba and Bb fragments [113]. Factor D, a trace plasma protein, is a serine protease without a zymogen form (i.e., it is always functionally active) [66]. Factor B is both structurally and functionally homologous to C2 of the classical pathway [114]. Like C2, factor B is a serine protease that exists in its native state as a zymogen with enzymatic cleavage being required for activation [115]. Further, activated C2 (C2b) and factor B (Bb) have the same natural substrates (i.e., C3 and C5). Although factor B and C2 share only modest (37%) amino acid sequence homology, the genes that encode these two proteins are

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structurally similar and closely associated in the MHC region of chromosome 6. Together, these observations suggest that C2 and factor B arose by gene duplication. Activation of the alternative pathway on erythrocytes is initiated when native C3 binds to C3 ⭈ H2O:Bb. The subsequent enzymatic cleavage of C3a from native C3 induces a conformational change in the resulting C3b molecule such that the internal thioester becomes exposed. By acyl transfer, nascent C3b binds covalently to glycophorin A, the major erythrocyte sialoglycoprotein [116]. Although both ester and imidoester bonds can form, the majority of C3b on erythrocytes are bound via an ester bond to the carbohydrate moiety of glycophorin A [117]. The cell surface C3 convertase is formed when factor B binds to C3b in a magnesium-dependent reaction (Fig. 3-1). Factor B is activated by factor D [113], and the amplification C3 convertase (C3bBb) is stabilized by factor P (properdin) [118] (Fig. 3-1). Both plasma factors and membrane proteins control the formation and stability of the alternative pathway C3 convertase (Fig. 3-1). Factor H [119] regulates the formation and stability of the convertase in three ways: first, by binding to C3b, it inhibits factor B binding; second, by binding to C3bBb, it displaces Bb from the complex; third, by binding to C3b, it acts as a cofactor for the enzymatic degradation of C3b to iC3b by factor I [68, 120, 121]. DAF (CD55) is the erythrocyte membrane constituent that is primarily responsible for regulating the activity of the alternative pathway C3 convertase. It binds to the C3bBb complex, causing the enzymatic subunit to dissociate [56, 57]. CR1 also controls C3 convertase activity. Using isolated complement components, purified CR1 has been shown to accelerate the decay of the alternative pathway C3 convertase in vitro [122]. In vivo, however, it appears likely that erythrocyte CR1 functions primarily as a factor 1 cofactor rather than as a regulator of the C3 convertase; blocking CR1 function does not enhance alternative pathway C3 convertase activity when serum is used as the complement source [123]. Available evidence suggests that CR1 exerts its activity intercellularly (binding to C3b on neighboring cells), while DAF functions as an intracellular regulator of the convertase (binding to C3bBb on the same cell) [124]. The C5 convertase of the alternative pathway consists of C3bBbC3b; one of the C3b molecules acts as the binding site for Bb and the other molecule serves to position C5 for cleavage by Bb [125] (Fig. 3-1). As is the case for the C3 convertase, the C5 convertase is stabilized by factor P, and the same plasma and membrane constituents that regulate the C3 convertase/control the activity of the C5 convertase (Fig. 3-1). Generation of C5b as a result of enzymatic cleavage of C5 by Bb releases C5a and initiates formation of the MAC (Fig. 3-1) in a manner analogous to that described above for the classical pathway. As is the case for the classical pathway, the cytolytic activity of the MAC formed on erythrocytes by the alternative pathway is also inhibited by MIRL (CD59) (Fig. 3-1). As discussed above, hemolysis of PNH erythrocytes in acidified serum is mediated by the alternative pathway. Compared to neutral pH, initiation and amplification of the alternative pathway C3 and C5 convertases are enhanced at pH 6.4 [125a]. Thus, the greater activity of the alternative pathway under

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slightly acidic conditions explains the observations of Stru¨bing, Hijmans van den Berg, and particularly Ham. A characteristic feature of processes mediated by the alternative pathway is that, in contrast with classical pathway-mediated lysis, the lytic event is not observed following modest dilution of serum (1 : 4 or 1 : 8). This explains why early investigators dismissed complement as the mediator of lysis of PNH erythrocytes. We now know that the role of properdin in the alternative pathway is different from that envisioned by Pillemer [17] who thought it functioned like antibody in the classical pathway. The alternative pathway does not require an initiating factor, however, and the function of properdin is to stabilize and thereby extend the enzymatic half-life of the amplification C3 (C3bBbP) and C5 convertases (C3bBbC3bP). Nonetheless, Pillemer’s vision of the properdin pathway as the effector of innate immunity proved prescient. The discovery that the properdin system mediates lysis of PNH erythrocytes in acidified serum solved a problem that had challenged many early investigations of PNH.

Phenotypic Mosaicism In discussing the acidified serum lysis test, Dacie [20] commented in his 1963 review that ‘‘Even if the serum is changed several times there appear to be always some cells which resist hemolysis.’’ PNH erythrocytes were abnormally sensitive to lysis when complement was activated by antibody, but complete hemolysis was never observed. These observations suggested the existence either of one population of cells that varied in sensitivity to lysis or of two populations, one abnormally sensitive to lysis and the other of normal sensitivity. There is a biphasic red cell survival pattern for transfused PNH erythrocytes with an early, steep slope suggesting a population with a very short life span, and a subsequent, less steep slope approximating that observed for normal erythrocytes. Dacie [20] favored the hypothesis that the erythrocytes of PNH consisted of two populations. Compared to normal erythrocytes, he also hypothesized that the greater sensitivity of PNH erythrocytes to antibody-initiated lysis was due to aberrant interactions with complement [20]. In the early 1960s, the work of Manfred Mayer made possible quantitative analysis of complement activity. Working with Dacie, Rosse modified his technique to assay serum complement in order to analyze the complement sensitivity of PNH erythrocytes [126], and he could demonstrate the magnitude of the difference in sensitivity between PNH and normal erythrocytes and clearly separate PNH erythrocytes into two quantitatively definable populations. In the complement lysis sensitivity (CLS) assay of Rosse and Dacie [126], erythrocytes were incubated with an excess of sensitizing antibody and with incremental concentrations of serum as the complement source; lysis was quantitated based on release of hemoglobin as determined spectrophotometrically. When the results were plotted as log (fraction of cells lysed/fraction of cells unlysed) against log complement concentration, a straight line was observed for

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normal erythrocytes [126] (Fig. 3-3). In contrast, erythrocytes from patients with PNH showed two connected, nearly parallel, straight-line portions of the curve (Fig. 3-3). These findings were interpreted to mean that the peripheral blood of patients with PNH consisted of two populations of cells that differed in susceptibility to complement-mediated lysis. Because the insensitive population was somewhat more susceptible to complement than normal, however, they concluded that ‘‘PNH cannot simply be regarded as a mosaic of abnormal cells proliferating along with normal cells but, in most cases, as a mosaic of two abnormal populations.’’ The results established unequivocally that abnormal sensitivity to complement underlies the greater hemolysis of PNH erythrocytes ‘‘regardless of the factors responsible for the initiation of the immune reaction,’’ a conclusion strongly supported by the observation that the cells that underwent

10

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Lysis with anti-I

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Complementinsensitive cells

Normal cells

1.0 Complementsensitive cells

0.1

0.01 1

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Figure 3-3 Demonstration of complement-sensitive and complement-insensitive populations in a patient with PNH using the complement lysis sensitivity assay. RBC from a patient with PNH (䊉) and from a normal volunteer (triangles) were incubated with antibody and incremental dilutions of serum as the complement source; hemolysis was subsequently quantitated. The antibody was in excess and the complement concentration was the limiting factor. The inflection point of the curve that marks the end of lysis of the complement-sensitive cells and the beginning of lysis of the complement-insensitive cells is indicated on the graph by the solid lines. The dashed lines mark the dilution of serum required to produce 50% lysis of the normal RBC and of the two populations of PNH cells. [Modified from Rosse, W. F. (1973). Variations in the red cells in paroxysmal nocturnal hemoglobinuria. Br. J. Haematol. 24, 327–342, by copyright permission of Blackwell Science Ltd.]

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hemolysis in the acidified serum lysis test were the same sensitive population defined by the complement lysis sensitivity assay [126]. Finally, by examining multiple patients, variation in the proportion of complement-sensitive cells was also demonstrated (ranging was from 4 to 80% in 11 cases) [126], providing a plausible explanation for the clinical variability in the severity of the hemolytic component of the disease. The studies of Rosse and Dacie represented a major conceptual advance and defined many of the fundamental characteristics of PNH. In particular, the recognition that the basic defect underlying the disease was represented in the complement-sensitive population provided a paradigm: if a process were specific for PNH erythrocytes, it should be observed in the complement-sensitive population but not in the insensitive population. By separating cell populations into

Fraction lysed

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10.0

1.0

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Figure 3-4 Demonstration of three different RBC phenotypes in a patient with PNH. Erythrocytes from two patients with PNH and a normal volunteer (triangles) were analyzed by using the complement lysis sensitivity assay. The RBC from one of the patients with PNH (䊊) consisted of two populations, a complement-sensitive and a complement-insensitive population. The RBC from the other patient (䊉) consisted of the following three populations: a markedly sensitive population (PNH type III); a population of intermediate sensitivity (PNH type II); and a population with nearly normal sensitivity (PNH type I). The dilution of serum required to produce 50% lysis of each PNH type from that patient is indicated by the dashed line. [Modified from Rosse, W. F. (1973). Variations in the red cells in paroxysmal nocturnal hemoglobinuria. Br. J. Haematol. 24, 327–342, by copyright permission of Blackwell Science Ltd.]

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complement-sensitive and complement-insensitive groups, the importance of the deficiency of erythrocyte acetylcholinesterase in PNH [20, 127, 128] was firmly established [129]—which 25 years later was related to the GPI-anchored nature of the protein [130]). Studies indicating that the complement-sensitive erythrocytes were monoclonal while the insensitive cells were polyclonal [131] proved critical for the hypothesis that somatic mutations account for the phenotypic mosaicism characteristic of PNH [20]. In 1969, Aster and Enright [132] demonstrated that platelets and neutrophils from PNH patients were also abnormally sensitive to complement-mediated lysis. The somatic mutation must occur in a primitive hematopoietic stem cell. By using a modification of the complement lysis sensitivity assay. Stern and Rosse [133] confirmed that like the erythrocytes, the granulocytes of PNH are also a mosaic. As noted above, the red cells of patients with PNH were initially divided into two groups (sensitive and insensitive) based on susceptibility to hemolysis by complement [126]. According to this classification, the insensitive population was on average approximately twice as sensitive to complement-induced hemolysis as normal erythrocytes. However, with more experience using the assay, it became apparent that the complement-insensitive population was heterogeneous [134] and three populations of erythrocytes could be identified in some patients with PNH (Fig. 3-4). Rosse [135] proposed a phenotypic classification shown in Table 3-3. In the vast majority of patients, a population of cells with complement sensitivity equivalent to that of normal cells was observed, along with a variable proportion of abnormally sensitive cells. Erythrocytes of intermediate sensitivity (the PNH II phenotype) could be identified coexisting with cells of normal sensitivity (the PNH I phenotype), marked sensitivity (the PNH III phenotype), or both [134–137]. The existence of three different red cell phenotypes challenged the hypothesis that PNH arose by simple monoclonal expansion [134]. The functional basis of the variability in complement sensitivity of PNH erythrocytes

TABLE 3-3 PNH Phenotypes a

Phenotypic designation PNH I PNH II PNH III

a

Complement sensitivity b Normal Moderately sensitive (3 to 4 times more sensitive than normal) Markedly sensitive (15 to 20 times more sensitive than normal)

GPI-AP expression by flow cytometry c Normal Dim positive Negative

Type of PIG-A mutation None Missense (partial inactivation of PIG-A) Nonsense, frameshifts deletions, insertions (complete inactivation of PIG-A)

Modified from Parker, C. J., and Lee, G. R. (1998). Paroxysmal nocturnal hemoglobinuria. In ‘‘Wintrobe’s Clinical Hematology’’ (G. R. Lee, J. Foerster, J. Lukens, F. Paraskevas, J. P. Greer, and G. M. Rodgers, Eds.), 10th Ed., pp. 1264–1288. Williams & Wilkins, Philadelphia, by copyright permission of Williams & Wilkins. b Based on the complement lysis sensitivity assay of Rosse and Dacie [11] (illustrated in Figs. 3-3 and 3-4). c Based on flow cytometry of erythrocytes (illustrated in Figs. 3-13 and 3-14).

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was delineated in 1989 [138], and in 1996, the molecular basis of the phenotypic mosaicism was defined [139]. The pathophysiological pressure that selects for the abnormal stem cells, however, remains enigmatic. Any hypothesis that attempts to explain PNH must account for this remarkable feature of phenotypic mosaicism that was defined originally by careful analysis of complement sensitivity of the erythrocytes.

Functional Basis of the Abnormal Sensitivity of the Erythrocytes of PNH to Complement-Mediated Lysis In 1973, Logue et al. [23] reported that PNH erythrocytes bound more C3 than normal erythrocytes when complement was activated by either the classical or the alternative pathway; for a given amount of C3 bound, PNH erythrocytes lysed to a much greater degree than normal cells. Apparently both quantitative and qualitative differences in complement interactions with PNH erythrocytes caused greater lytic sensitivity. By analyzing the relationship between the extent of lysis and the amount of C3 bound, Rosse and colleagues [135] demonstrated differences between PNH II and PNH III erythrocytes. They found that while both phenotypes bound equivalent, supranormal amounts of C3, the type III cells underwent more lysis per C3 molecule bound when compared with type II cells. These results suggested that either PNH III cells had an additional defect that was not shared by PNH II cells, or that the abnormality was the same for both types but that the type III cells were more severely affected than the type II cells. Using an indirect experimental approach, Rouault and colleagues [140] found that, compared to normal erythrocytes, the MAC was more efficient at inducing lysis of PNH III cells. Hemolysis was induced by the classical pathway, and by varying the concentration of antibody, conditions were chosen so that normal and PNH III cells lysed to the same extent; the number of MAC lesions formed was counted using electron microscopy. For the same extent of lysis, approximately 10 times more MAC lesions were observed on normal erythrocytes compared to PNH III erythrocytes. Greater susceptibility of PNH III cells to complement-mediated lysis was due, at least in part, to the greater efficiency of the MAC when activated on PNH cells. The efficacy of the MAC on PNH erythrocytes was also investigated by Packman et al. [141], who demonstrated that type III cells were abnormally susceptible to reactive lysis but type II cells were not. In reactive lysis systems (also known as passive or bystander lysis) [142, 143], the entire complement cascade is not activated directly on the surface of the erythrocyte, as it is when hemolysis of PNH cells is induced either by the classical or alternative pathways. Rather, the C3 convertase step is bypassed in reactive lysis systems, the MAC is directly generated on the cell surface (Fig. 3-5) and the effects of the MAC are directly examined. These studies provided the first direct evidence in support of the greater lytic susceptibility of PNH III erythrocytes compared to PNH II cells as a result of the greater efficiency of the MAC-induced lysis of the type III cells. The basis of this difference between the type II and III cells and the nature of the aberrant effects of the MAC on type III cells, however, remained enigmatic.

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Figure 3-5 Reactive lysis. In reactive lysis, complement is not directly activated on the cell surface. Rather isolated C5b6 complexes are added to buffer containing the indicator cell (A). The C5b6 complexes are hydrophilic and do not bind to the membrane, but when isolated C7 is added to the reaction mixture, it binds to the bimolecular complex. Consequently, a comformational change is induced, resulting in an amphilic transition that exposes a labile membrane binding site in the C7 molecule (indicated by the star) (B). If the C5b67* complex does not encounter a cell, the binding site is rapidly lost as a result of spontaneous inactivation. If the complex encounters an RBC, the C7 component inserts into the membrane via hydrophobic interactions. Following the addition of a source of C8 and C9 (either isolated components or serum), a single molecule of C8 binds to the C5b component of the trimolecular complex and inserts into the lipid bilayer (C). The membrane attack complex (MAC) is formed when multiple molecules of C9 (C9n) bind spontaneously to the complex (D). The C9 molecules polymerize forming a membrane channel, and the cell consequently undergoes colloid osmotic lysis.

Identification of Erythrocyte Membrane Proteins That Regulate the C3 Convertase Both PNH II and PNH III erythrocytes bound more C3b than normal erythrocytes when complement was activated by either the classical or the alternative pathway [23, 135, 141]. These results suggested that regulation of the C3 convertases of both pathways might be aberrant on PNH erythrocytes. During the 1970s, methods for purifying components of the complement cascade became available, and by 1980, it was possible to assemble the entire C3 convertase of the alternative pathway using isolated components. The utilization of purified components allowed a more precise view of the influence of the cell surface on the assembly and stability of the convertase complex. Using isolated components of the alternative pathway C3 convertase, Parker et al. [144] and Pangburn et al. [145] demonstrated that the activity, formation, and stability of the convertase were enhanced on PNH cells, providing a clear explanation for the greater binding of activated C3 but not a definition of the molecular basis of the abnormal convertase activity. In 1979, Fearon [122] reported the isolation of a protein from normal human

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erythrocytes that inhibited the activity of the C3 convertase of the alternative pathway: by binding to cell-associated C3b, this protein prevented the formation of the amplification convertase. This regulatory factor also accelerated the decay of formed convertases (C3bBb) by binding to C3b and thereby dissociating the enzymatic subunit (Bb). In addition, the isolated erythrocyte membrane protein was shown to act as a cofactor for factor I in the enzymatic degradation of C3b. Subsequent studies [146] demonstrated that the inhibitory factor was the membrane receptor for C3b known as complement receptor type I (CR1 or CD35). Functional properties of CR1 suggested that its absence from erythrocytes could result in greater binding of C3b to cells, as the formation and stability of the C3 convertase would be enhanced. The functional activity of CR1 on PNH erythrocytes was found to be normal [147], and blocking CR1 activity on normal erythrocytes by using specific antibodies failed to enhance the susceptibility of the cells to complement-mediated lysis. However, PNH erythrocytes expressed subnormal amounts of CR1 [117, 148], a deficiency, that appeared to be an epiphenomenon resulting from exposure of the erythrocytes to complement activation in vivo and not specific for PNH cells [117, 148]. Although CR1 was found not to be involved in the pathophysiology of PNH, isolation and characterization of the protein was nonetheless important because it confirmed the existence of discrete membrane constituents that function specifically as complement regulators. Further, Fearon’s studies demonstrated that complement regulatory factors that are integral membrane proteins could be purified to homogeneity while retaining functional activity. A decade earlier (1969), Hoffmann [149, 150] had published results of a series of experiments showing that an n-butanol extract prepared from human erythrocyte stroma contained a factor or factors that inhibited complementmediated hemolysis. A portion of the active material possessed the capacity to bind to the indicator cell and remain functionally active. Subsequently, the name decay-accelerating factor (DAF) was applied to the functional property of the extract because the material enhanced the rate at which the activity of the classical pathway C3 convertase diminished over time [151, 152]. Based on these observations, Chua et al. [153] hypothesized that deficiency of DAF could account for the greater fixation of C3 to PNH erythrocytes. The n-butanol extract from normal erythrocytes was found to be a potent inhibitor of complement-mediated lysis of PNH erythrocytes. When either the DAF activity or the complement lysis inhibitory activity of erythrocyte extracts derived from PNH and normal cells was compared, however, no difference was observed. Even the extract from uniform populations of PNH III erythrocytes was found to have normal complement lysis inhibitory activity. Technical issues likely accounted for the negative results, as Chua et al. used detergent rather than nbutanol to prepare some of the extract, and under these conditions, CR1 would have been present in the preparations. Thus, solubilized CR1 may have been responsible for the decay-accelerating activity observed in the PNH III cells.) One of the complement-regulatory proteins contained in the n-butanol extracts described by Hoffmann was purified to homogeneity by Nicholson-Weller and colleagues [55]: This protein was named decay-accelerating factor of stroma (DAF-S). The term DAF-S was introduced to distinguish the membrane-derived from serum-derived factors with decay-accelerating activity that were being char-

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acterized at the same time by others. (The DAF-S designation did not endure, however, and the protein isolated by Nicholson-Weller and colleagues quickly became known simply as DAF.) The serum-derived DAFs were named individually (i.e., C4b binding protein and factor H) after they were isolated and characterized. In addition to its capacity to restrict the activity of the classical pathway C3 convertase, DAF was also found to inhibit the activity of the alterantive pathway C3 convertase [55]. The functional properties of DAF strongly suggested that its absence from erythrocytes would lead to greater C3 convertase activity and hence to greater C3b deposition on the cell surface, regardless of whether complement was activated by the classical or by the alternative pathways. Absence of DAF could therefore account for the observed differences between PNH and normal erythrocytes (Fig. 3-6). That PNH erythrocytes were deficient in DAF was reported by Nicholson-Weller and colleagues in 1983 [154]; PNH II cells were partially deficient while PNH III cells were completely DAF deficient. Almost simultaneously, Pangburn et al. [155] presented both functional and immunochemical evidence of DAF deficiency in PNH. The discovery of this

Figure 3-6 DAF inhibits the C3 convertase on normal human erythrocytes. DAF regulates the alternative pathway C3 convertase by binding to the bimolecular C3bBb complex (A). This binding dissociates the enzymatic subunit of the convertase (Bb*), rendering the complex inactive (B). DAF has a low affinity for C3b. Thus, once Bb dissociates, DAF is free to bind to another C3 convertase. The deficiency of DAF on the erythrocytes of PNH enhances the stability of the C3 convertase (C). This increased stability results in greater C3 convertase activity, resulting in greater binding of C3 to PNH cells (D). On human erythrocytes, activated C3 binds primarily to glycophorin A via ester bond formation (-o-) with the carbohydrate moiety (䊉) of this abundant sialoglycoprotein. DAF regulates the C3 convertase of the classical pathway in a manner analogous to that of the alternative pathway. In this case, DAF binds to the C4bC2b complex and dissociates C2b, the enzymatic subunit of the classical pathway convertase (Fig. 3-1).

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deficiency was a major milestone in the journey toward understand the basis of the hemolysis of PNH that began with the observations of Stru¨bing, Hijmans van den Berg, and Ham. Subsequently Medof and colleagues [156] showed that by using monoclonal antibodies to block DAF function, normal erythrocytes could be made sensitive to hemolysis in acidified serum. Further, they demonstrated that incubation of PNH erythrocytes with purified DAF ameliorated susceptibility to lysis in acidified serum. These experiments established a causal relationship between the deficiency of DAF and the aberrant interactions of PNH erythrocytes with complement. DAF is a 70-kDa single chain glycoprotein (reviewed in [56, 57]). As noted above, DAF contains four SCR units that are homologous to domains found in other complement-regulatory proteins (e.g., CR1, CR2, C4b binding protein, factor H, and membrane cofactor protein). The DAF gene maps to chromosome 1 (band q32) and forms part of the regulators of complement activation (RCA) locus that includes the genes that encode CR1, CR2, C4b binding protein, factor H, and membrane cofactor protein. DAF destabilizes the C3 convertases both of the classical and alternative pathways; it does not inhibit either binding of C2 to C4b or binding of factor B to C3b. Rather, once the convertase is formed (Fig. 3-1), DAF causes the catalytic subunit (C2b in the case of the classical pathway and Bb in the case of the alterantive pathway) to dissociate [157]. The deficiency of DAF provided a logical explanation for the greater binding of C3b to PNH erythrocytes when complement was activated either by antibody (classical pathway) or by acidification of serum (alternative pathway) and a causal role for DAF in the pathophysiology of PNH was readily accepted. But could DAF deficiency explain all of the aberrant interactions of complement with PNH erythrocytes? As noted above, compelling evidence had indicated that regulation of the MAC was abnormal on PNH erythrocytes [96, 140, 141]. Thus, if DAF deficiency alone were sufficient to account for the lytic sensitivity of PNH erythrocytes, DAF would have to play a role (direct or indirect) in the regulation of the MAC. However, DAF had no inhibitory activity in the process of reactive lysis [158, 159], and by implication another protein that functioned as a regulator of the MAC must also be deficient in PNH. That more than one protein was deficient in PNH was supported by previous work that had shown that erythrocyte acetylcholinesterase was also deficient in PNH, and acetylcholinesterase levels correlated with the proportion of complement-sensitive cells [127–129]. Earlier studies had also shown that neutrophils from patients with PNH were similarly low in alkaline phosphatase [160, 161]. The probability that discrete mutations affecting different gene proteins individually accounted for the abnormality seemed unlikely. Conceivably, a large deletion could effect the loss of multiple proteins, if the genes were in relatively close proximity on the same chromosome, but arguing against this mechanism was the absence of consistent karyotypic abnormalities in PNH. A more probable explanation for the deficiency of multiple proteins was that the affected membrane constituents shared a common posttranslational modification, and that some step in the processing of this moiety was abnormal. A tenet of this hypothesis was that the genes that encode the deficient proteins in PNH would be structurally and functionally normal. As anticipated, neither mutations nor abnormalities in DAF gene expression are found in PNH [162]. Meanwhile, the post-

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translational modification (the GPI anchor) that is shared by all of the proteins that are deficient in PNH was identified in 1986 [130, 163].

Identification of Erythrocyte Membrane Proteins That Inhibit the MAC As DAF deficiency could not account for abnormal regulation of the MAC on PNH erythrocytes, attention was focused on identifying an erythrocyte membrane protein that inhibited the lytic activity of the terminal complement complex (C5b-9). Using a reactive lysis system in which whole serum complement was activated by cobra venom factor (CVF) (Fig. 3-7), Parker et al. [96] showed that PNH III erythrocytes bound much greater amounts of C9 than either PNH II or normal cells, findings that confirmed that PNH erythrocytes lacked a membrane factor to regulate assembly of the MAC. A short time later, a protein that regulated susceptibility to reactive lysis was isolated from normal red cells, termed C8-binding protein (C8bp) by Scho¨nermark et al. C104 and homologous restriction factor (HRF) by Zalman and colleagues [165]. The factor blocked formation of the MAC by binding to C8, C9, or both. In 1987, each group

Figure 3-7 Cobra venom factor (CVF) initiated lysis. CVF is a structural analog of human C3c, but it has the functional properties of human C3b. In the presence of MgCl2 (:), CVF binds factor B. Factor D cleaves a portion (the Ba fragment) of CVF-bound factor B, generating the enzymatic subunit of the complex (Bb). When CVF:Bb complexes are added to whole serum, C5 binds to CVF portion of the CVF:Bb complex, positioning it for cleavage. Consequently, the C5a fragment is excised enzymatically by Bb, and the resulting nascent C5b molecule forms a stable, hydrophilic complex with C6 (A). The C5b6 complex does not directly attach to the membrane, but when C7 binds to the bimolecular complex, a comformational change is induced. As a result, an amphilic transition occurs that exposes a labile membrane binding site in the C7 molecule (indicated by the star) (B). If the C5b67* complex does not encounter a cell, it is rapidly inactivated as a result of interaction with endogenous plasma inhibitors such as vitronectin (VN) or clusterin (CL) (B). If the complex encounters an RBC, the C7 component inserts into the membrane via hydrophobic interactions. Next, C8 (C) and multiple molecules of C9 (C9n) (D) bind spontaneously and in sequence, thereby forming the cytolytic membrane attack complex (MAC).

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reported that C8bp/HRF was deficient in PNH [166, 167]. The basis of the aberrant interaction of the MAC on PNH erythrocytes appeared to have been identified. The role of C8bp/HRF was called into question by the isolation, in 1989, of a protein from human erythrocytes that inhibited CVF-initiated lysis of PNH erythrocytes, which Holguin et al. [105] named membrane inhibitor of reactive lysis (MIRL). MIRL seemed discrete from C8bp/HRF because the relative mass of the two proteins (18 kDa for MIRL and 64 kDa for C8bp/HRF) was very different, and immunochemical studies indicated that anti-MIRL antibodies reacted with only one erythrocyte membrane protein (Fig. 3-8). MIRL was also

Figure 3-8 Analysis by western blot of normal and PNH III erythrocyte membrane proteins using anti-MIRL as the primary antibody. Aliquots of 5 애g of hemoglobin-free erythrocyte ghosts were subjected to SDS-PAGE under nonreducing conditions and electrophoretically transferred to nitrocellulose paper. After incubation with anti-MIRL antiserum, antibody binding was localized by using alkaline phosphatase-conjugated anti-rabbit IgG and a chromogenic substrate. The PNH III erythrocytes are deficient in MIRL. [Reproduced form Holguin, M. H., Fredrick, L. R., Bernshaw, N. J., Wilcox, L. A., and Parker, C. J. (1989). Isolation and characterization of a membrane protein from normal human erythrocytes that inhibit reactive lysis of the erythrocytes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 7–17, by copyright permission of the American Society of Clinical Investigation.]

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shown to be deficient on PNH III erythrocytes (Fig. 3-8). Further, MIRL could incorporate into the membrane of PNH III erythrocytes and protect them from reactive lysis (Fig. 3-9). This capacity suggested that, like other proteins that are deficient in PNH, MIRL was GPI anchored. Finally, by blocking MIRL function with antibody, normal erythrocytes could be made susceptible to reactive lysis (Fig. 3-10). These data provided compelling evidence that MIRL deficiency was an important component of the pathophysiology of PNH (Fig. 3-11) and cast doubt upon the importance of C8bp/HRF as a regulator of the MAC. Whether deficiency of C8bp/HRF contributes to the abnormal susceptibility of PNH erythrocytes to complement-mediated lysis is doubtful. When MIRL function is blocked with monospecific antibody, normal erythrocytes are made as susceptible to reactive lysis as PNH III erythrocytes despite normal C8bp/ HRF function (MIRL antibodies do not cross-react with any other erythrocyte membrane constituent). Additionally, there are no data on the primary structure of C8bp/HRF nor has a candidate cDNA or gene been identified despite the fact that the protein was discovered 13 years ago—thus, the very existence of C8bp/HRF is in doubt.

Figure 3-9 MIRL inhibits cobra venom factor initiated lysis of PNH III erythrocytes. Normal erythrocytes (䊉) were incubated with buffer and PNH erythrocytes were incubated with buffer (shaded triangles) or with buffer containing purified MIRL (5 애g/ml) (open triangles). After washing, the cells were incubated with cobra venom factor complexes (CVF:Bb) (Fig. 3-7) and incremental concentrations of human serum as the complement source. MIRL that incorporated into the membrane-protected PNH III RBC against cobra venom factor initiated lysis. [Reproduced from Holguin, M. H., Fredrick, L. R., Bernshaw, N. J., Wilcox, L. A., and Parker, C. J. (1989). Isolation and characterization of a membrane protein from normal human erythrocytes that inhibits reactive lysis of the erythrocytes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 7–17, by copyright permission of the American Society of Clinical Investigation.]

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Figure 3-10 Normal erythrocytes are rendered susceptible to cobra venom factor initiated lysis after treatment with anti-MIRL. Normal erythrocytes were incubated with incremental concentrations of anti-MIRL antiserum (䊉) or with nonimmune rabbit serum (shaded triangles). After washing, the cells were incubated with CVF:Bb complexes (Fig. 3-7) and serum (the complement source). Hemolysis was subsequently quantitated. Inhibition of MIRL function with antibody, induced susceptibility to cobra venom factor initiated lysis. [Reproduced from Holguin, M. H., Fredrick, L. R., Bernshaw, N. J., Wilcox, L. A., and Parker, C. J. (1989). Isolation and characterization of a membrane protein from normal human erythrocytes that inhibits reactive lysis of the erythrocytes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 7–17, by copyright permission of the American Society of Clinical Investigation.]

Figure 3-11 Inhibition of the MAC by MIRL. MIRL (CD59) inhibits MAC formation in two ways. First by binding to C8 within the C5b-8 complex, it inhibits the spontaneous binding of C9 (A). Second, by binding to C9 within the C5b-9 complex, MIRL blocks C9 multiplicity, thereby inhibiting the polymerization of C9 that is needed for generation of a cytolytic MAC (B). PNH RBC lack MIRL. Consequently, MAC formation proceeds unimpeded, and the cell undergoes complement-mediated lysis (C and D).

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In contrast, the structure and function of MIRL (CD59) have been extensively characterized. The protein was independently identified by several groups [169–174], although none of those investigators were attempting to identify the factor that accounted for the abnormal sensitivity of PNH erythrocytes to reactive lysis. Different names have been proposed for the protein: CD59, protectin, MACIF, HRF 20, MEM-43, and H19 (the term MIRL is used here, because that was the name given to the factor that was absent from PNH III erythrocytes and accounted for their abnormal sensitivity to MAC-induced lysis). MIRL is an 18-kDa GPI-anchored protein with wide tissue distribution ([173, 174] and reviewed in [106]). It inhibits the lytic activity of the MAC primarily by binding to the C8 component of the C5b-8 complex and to the C9 component of the C5b-9 complex (Fig. 3-11). Binding to C5b-8 appears to inhibit C9 binding, and binding to C9 within the C5b-9 complex appears to restrict C9 multiplicity [106–109]. Normal human erythrocytes express 25,000–30,000 copies of MIRL/RBC (there are approximately 10 times more copies of MIRL than DAF per RBC). In the case of DAF, determination of the primary sequence provided insight into the functional properties of the protein because DAF was found to share a common structural motif (the SCR unit) with other proteins that bind to activated C3 and C4 (see discussion above). For MIRL, however, delineation of the primary structure revealed no significant homology with any other complement-regulatory factors. MIRL shares modest homology with a number of other proteins, including members of the murine Ly-6 multigene family, a squid protein, the receptor for urokinase plasminogen activator, and snake 움-neurotoxins [170, 171, 175–180]. The common feature that unifies these functionally diverse proteins is the conservation of Cys residues. MIRL is a prototypic example, containing 10 Cys that are paired to form five intrachain disulphide bonds [181]. As a result of these disulphides, the locked loop structures that are characteristic of this group of proteins are generated. By NMR, the three-dimensional structure of MIRL has been determined. [182–184] The molecule appears disc shaped and flat and it consists of two antiparallel 웁-sheet regions, a short helical region, and a carboxy-terminal region devoid of secondary structure. MIRL contains a single, large N-linked carbohydrate moiety that accounts for approximately one-third of its molecular mass. The structure of the N-linked sugar is heterogeneous and complex, but carbohydrate residues are not required for the MAC regulatory activity of MIRL [185–188]. The gene that encodes MIRL [177, 189] is located on the short arm of chromosome 11 (p14–p13).

Molecular Basis of the Erythrocyte Phenotypes of PNH As previously discussed PNH III erythrocytes had been shown to be susceptible to reactive lysis, whereas PNH II erythrocytes were resistant to this process. Either of two hypotheses seemed plausible as explanations for the molecular basis of the differences between the PNH II and the PNH III phenotypes. According to the first, the two phenotypes have the same basic defect but PNH III cells are more severely affected. According to the second hypothesis, PNH III erythrocytes have two independent defects: DAF deficiency would account for the intermediate complement sensitivity (resulting from the greater activity

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of the C3/C5 convertase) defect shared by both PNH II and PNH III cells; the further absence of MIRL would be responsible for the greater susceptibility of PNH III cells to reactive lysis. Compelling data showed that the first hypothesis is correct [138]. Using immunochemical techniques, DAF and MIRL were quantitated on PNH II and PNH III erythrocytes [138]. As anticipated, the PNH III cells were almost completely deficient in both regulatory proteins (Fig. 3-12). PNH II erythrocytes were also shown to be markedly deficient in both regulatory proteins, but the deficiency was less severe in comparison with PNH III cells (Fig. 3-12). These results indicated that erythrocytes that are classified as PNH II have an amount of MIRL that is abnormally low but still above the threshold that provides protection against reactive lysis. On PNH erythrocytes, the deficiency of DAF and MIRL are concordant. In general, PNH II cells have approximately 10% of the amount of DAF and MIRL expressed by normal erythrocytes, PNH II cells express about 250–300 copies of DAF and about 2500–3000 copies of MIRL. This amount of MIRL is sufficient to inhibit the lytic activity of the MAC in reactive lysis systems [105], but the level of DAF is insufficient to control the activity of the C3 convertase when complement is activated by either the classical or the alternative pathway. Currently, erythrocyte phenotypes of PNH are defined by flow cytometric analysis using anti-DAF and anti-MIRL as primary antibodies [190–194]; this technique is informative in its separation of different phenotypes and in illustration of the mosaicism characteristic of PNH (Figs. 3-13 and 3-14).

Figure 3-12 Quantitation of erythrocyte DAF and MIRL using a radioimmunobinding assay. Normal (PR) and PNH erythrocytes from patient RO (predominately type III cells), patient BA (predominately type II cells), and OB (a mixture of type I, type II, and type III) were incubated with a saturating amount of either anti-DAF or anti-MIRL. Subsequently, binding of rabbit IgG was quantitated following incubation with a saturating amount of radiolabeled anti-rabbit IgG. The values (specific binding) depicted by the bars represent the mean ⫹ SD (n ⫽ 5). There is a concordant deficiency of DAF and MIRL in both the PNH III and the PNH II phenotypes. Relatively subtle quantitative differences in expression of DAF and MIRL underlie the variability in complement sensitivity of PNH erythrocytes and thereby determine the phenotype. [Reproduced from Holguin, M. H., Wilcox, L. A., Bernshaw, N. J., Rosse, W. F., and Parker, C. J. (1989). Relationship between the membrane inhibitor of reactive lysis and the erythrocyte phenotypes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 1387–1394, by copyright permission of the American Society of Clinical Investigation.]

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Figure 3-13 Analysis of expression of DAF and MIRL on PNH erythrocytes. Erythrocytes from a patient with PNH and a normal volunteer (NL) were analyzed by flow cytometry using monoclonal antibodies. The histograms of the cells from the normal donor show uniformly positive staining with both antibodies. In contrast, the patient’s histograms demonstrate two discrete populations of cells. Approximately 30% of the cells are negative for DAF and MIRL (PNH III phenotype), and 70% show normal expression (PNH I phenotype). [Modified from Parker, C. J., and Lee, G. R. (1998). Paroxysmal nocturnal hemoglobinuria. In ‘‘Wintrobe’s Clinical Hematology’’ (G. R. Lee, J. Foerster, J. Lukens, F. Paraskevas, J. P. Greer, and G. M. Rodgers, Eds.), 10th ed., pp, 1264–1288. Williams & Wilkins, Philadelphia, by copyright permission of Williams Wilkins.]

Isolated Deficiencies of DAF and MIRL The relative importance of DAF and MIRL in regulating susceptibility to complement-mediated lysis is clear for instances in which there is an isolated deficiency of either, but not both, of the two inhibitors. Antigens of the Cromer blood group complex are located on DAF, and there are rare cases of a null phenotype called Inab [195–200]. Inab erythrocytes are deficient in DAF, but unlike PNH erythrocytes, other GPI-anchored proteins (including MIRL) are expressed normally. Following incubation in acidified serum, Inab erythrocytes bound about 20 times more activated C3 than normal erythrocytes, but no lysis was observed [197]. When MIRL function was blocked with antibody, the Inab cells became completely susceptible to acidified serum

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Figure 3-14 Phenotypic mosaicism in PNH. Erythrocytes from a patient with PNH and from a normal volunteer (NL) were analyzed by flow cytometry using either anti-DAF or anti-MIRL as the primary antibody. The histograms of the erythrocytes from the normal donor show uniformly positive staining with both antibodies. The patient’s histogram is consistent with three discrete populations of cells (a negative population, PNH type III; a population with partial expression, PNH type II; a population with normal expression, PNH type I). Statistical analysis of the three groups of cells from the patient showed that the PNH III population contributed 14% to the total, the PNH II population contributed 75%, and the PNH I population contributed 11%. [Modified from Parker, C. J. (1996). Molecular basis of paroxysmal nocturnal hemoglobinuria. Stem Cells 14, 396–411, by copyright permission of AlphaMed Press.]

lysis [197]. Therefore, DAF has regulatory activity for the alternative pathway C3 convertase but MIRL is primarily responsible for controlling susceptibility to hemolysis in acidified serum. Similar results are observed when antibodies are used to block the function of DAF and MIRL separately and together (Fig. 3-15) [123]. The conclusions reached by in vitro experiments are supported by in vivo observations that individuals with the Inab phenotype do not have hemolytic anemia [200]. Activation and degradation products of C3 cannot be detected on fresh Inab cells, indicating that the alternative pathway C3 convertase is not activated on Inab cells in vivo [197]. In contrast to the situation of individuals with the Inab phenotype, a patient with an isolated, inherited deficiency of MIRL suffered from a clinical syndrome indistinguishable from that of PNH [110]. The patient, whose parents were cousins, was homozygous for a single base pair deletion, and, the resulting frameshift

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Figure 3-15 Molecular basis of acidified serum lysis. Normal human erythrocytes were incubated with a combination of anti-DAF and anti-MIRL (shaded triangles), anti-MIRL and nonimmune rabbit serum (open triangles), anti-DAF and nonimmune rabbit serum (shaded circles), or nonimmune rabbit serum (open circles). After washing, the cells were incubated in acidified serum and hemolysis was subsequently quantified. Blocking DAF and MIRL function caused normal cells to undergo hemolysis in acidified serum. Thus, absence of DAF and MIRL accounts for the sensitivity of PNH erythrocytes to lysis in acidified serum. [Modified from Wilcox, L. A., Ezzell, J. L., Bernshaw N. J., and Parker C. J. (1991). Molecular basis for the enhanced susceptibility of the erythrocytes of paroxysmal nocturnal hemoglobinuria to hemolysis in acidified serum. Blood 78, 820–829, by copyright permission of W. B. Saunders Company.]

introduced a premature termination codon [201]. He experienced recurrent episodes of intravascular hemolysis with hemoglobinuria, and during two hemolytic crises cerebral infarction was documented [110]. In vitro, both the acidified serum and sucrose lysis assays confirmed that the patient’s erythrocytes were abnormally sensitive to complement-induced lysis [202]. Thus, the PNH phenotype is primarily a manifestation of MIRL deficiency. Separating the contributions that DAF and MIRL deficiency make to the abnormal regulation of complement and to the hemolysis of PNH is somewhat artificial because both proteins are important regulators of complement. In PNH, it is the combined deficiency of both proteins that determines the phenotypic characteristic of affected cells [203]. In the event that it becomes practical to replace only one or the other protein (using gene therapy or by infusion of recombinant protein), in vitro studies suggest that restoration of MIRL activity will be more effective in controlling hemolytic manifestations of the disease.

Artificial PNH Cells Following the report of Sirchia and colleagues [204] in 1965, it became apparent that artificial PNH cells showing marked sensitivity to complement-mediated hemolysis could be obtained by treating normal human erythrocytes with 2amino-ethylisothiouronium bromide. Phenotypically, AET-treated cells resem-

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ble PNH III cells in their similar susceptibility in the complement lysis sensitivity assay and in acidified serum, and they are susceptible to cobra venom factor initiated hemolysis [23, 205–207]. Further, like PNH III cells, AET-treated cells bind relatively large amounts of C5b-9 during cobra venom factor initiated reactive lysis [96]. Unlike PNH III erythrocytes, however, AET-treated cells bind little more C3 than normal erythrocytes following activation of the classical pathway [23]. The susceptibility profile suggests that MIRL, but not DAF, is dysfunctional on AET-treated cells. In agreement, MIRL was not detected by Western blot under reducing conditions [105]. Further, AET-treated erythrocytes did not bind anti-MIRL antibody (Fig. 3-16) [138]. AET appears to destroy the structural and functional integrity of MIRL by disrupting disulphide bonds (as discussed above, all 10 of the Cys residues of MIRL are paired to form five separate intrachain disulphide bonds [181]). In contrast to MIRL, AET reduced to only about 50% the functional activity of DAF [207], which is surprising because Cys residues appear to be critically important constituents of the structural motif (the SCR unit) of most of the extracellular domain of DAF. Each SCR forms a protein loop held together by disulphide bonds derived from four conserved Cys residues [52]. Like DAF, CR1 is composed largely of SCRs, but in

100

Anti-MIRL

125

I-anti-rabbit-IgG bound (cpm-3)

Non immune 80

60

40

20

0 Buffer

AET

Buffer

Trypsin

Treatment Figure 3-16 Binding of anti-MIRL to AET and trypsin-treated erythrocytes. The binding of anti-MIRL to normal erythrocytes treated with AET, trypsin, or buffer was assessed using rabbit anti-MIRL antiserum as the source of the primary antibody and radiolabled anti-rabbit IgG as the secondary antibody. The values depicted by the bars represents the mean ⫹ SD (n ⫽ 5). Treatment with AET completely abolished the binding of anti-MIRL, whereas trypsin treatment had little effect. Disruption of the structural and functional integrity of complement-regulatory proteins accounts for the abnormal susceptibility of AET-treated erythrocytes to complement-mediated lysis. [Reproduced from Holguin, M. H., Wilcox, L. A., Bernshaw, N. J., Rosse, W. F., and Parker, C. J. (1989). Relationship between the membrane inhibitor of reactive lysis and the erythrocyte phenotypes of paroxysmal nocturnal hemoglobinuria. J. Clin. Invest. 84, 1387–1394, by copyright permission of the American Society of Clinical Investigation.]

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the case of CR1, the complement-regulatory activity of the protein is completely destroyed by AET [207]. The aberrant interactions of AET-treated cells with complement are due to partial (in the case of DAF or complete for MIRL and CR1) chemical inactivation of membrane proteins that regulate complement. Thus, the chemically modified cells differ from both PNH II and PNH III erythrocytes. Nonetheless, AETtreated cells can be useful models for analyzing the regulation of complement on human erythrocytes if these disparities with PNH are recognized in the interpretation of experimental results.

Treatment of Hemolysis Despite the great progres that has been made in understanding the molecular basis of the hemolysis of PNH, standard therapy remains empirical. Management must be individualized because the clinical manifestations of the disease are highly variable. Some patients have a compensated hemolytic process with only occasional hemoglobinuria. These mild cases may require no specific therapy, although they should receive supplemental folate (1 mg/day), and their iron stores should be monitored once or twice yearly. Approximately 15% of patients experience a spontaneous remission, and for patients living with the disease for greater than 10 years, as many as 34% remit spontaneously [208]. Patients with severe hemolysis require intervention. Currently, the only definitive therapy is marrow ablation followed by stem cell rescue (peripheral blood or bone marrow transplantation). Because of the associated morbidity and mortality, however, allogenic transplants have been reserved primarily for patients with either recurrent life-threatening thromboembolic disease or a hypocellular marrow resulting in clinically significant leukopenia, thrombocytopenia, or both. Patients with severe hemolytic anemia should not be eliminated from consideration of allogeneic transplantation, however. The chronic hemolysis of PNH can be debilitating, causing troubling constitutional symptoms (e.g., malaise, lethargy, asthenia), and exacerbations are associated with a variety of untoward, sometimes fatal consequences (e.g., dysphagia, abdominal pain, back pain, impotence, and thromboemoblic events). In the rare patient with an identical twin, strong consideration should be given to syngeneic transplant because the morbidity and mortality rates are very much lower compared to allogeneic transplantation, and long-term immunosuppressive therapy is not required. Prior to infusion of the donor marrow or peripheral blood stem cells, however, a conditioning regimen appears necessary [209, 210]: failure to ablate the marrow of the recipient almost invariably results in disease recurrence because the PIG-A mutant, GPI-AP⫺ stem cells appear to have a proliferative or survival advantage over normal stem cells [209]. In addition to eliminating the mutant stem cells, the conditioning regimen may also eliminate the as yet undefined pathological process that selects for the mutant cells. Corticosteroids can ameliorate the hemolysis of PNH, although mechanism of action is speculative. High doses of prednisone (in the range of 1 mg/kg/day) can be used to treat exacerbations. Responses are usually rapid (within 24 h of starting treatment). The dose can often be reduced after 7–10 days, and high

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dose, daily corticosteroids should not be continued for more than 3 weeks. To reduce the complications associated with chronic use of prednisone, an alternate day schedule is recommended. Doses of 15–40 mg qod are usually well tolerated, although careful monitoring for evidence of steroid-induced toxicity is essential. Therapy with androgenic steroids is reserved primarily for patients with inadequate hematopoiesis due to bone marrow failure, but the hemolysis of PNH may also respond. By stimulating erythropoiesis, androgens may enhance the capacity of the marrow to compensate for the hemolysis, rather than affect the hemolytic process directly. Paradoxically, if the newly stimulated hematopoiesis primarily involves the PIG-A mutant stem cell clones, hemolysis may worsen. Patients with inadequately compensated hemolysis who are unresponsive to corticosteroids or who require unacceptably high doses require transfusion. The common recommendation that blood be given in the form of saline-washed [211] or frozen/thawed, deglycerolized red cells [212] in order to avert a hemolytic episode has been questioned [213, 214]. ‘‘Hyper’’-transfusion to nearly normal hemoglobin concentrations can produce short-lived remission [211–215], presumably by suppression of erythropiesis and a reduction of production of abnormal erythrocytes by the mutant stem cells. Because the hemolysis of PNH is intravascular, much of the iron that is released into the plasma as a consequence of the complement-mediated lysis is lost in the urine [216–218]. Negative iron balance is observed in most patients with a significant hemolytic component, even though the patient may have received multiple transfusions. Generally, oral iron therapy is well tolerated, but hemolytic episodes have been precipitated by such treatment [217, 218]. (Stru¨bing [1, 3] reported that his patient experienced a severe exacerbation of hemoglobinuria after surreptitiously ingesting iron salts.) Hemolytic episodes have also been reported after administration of iron dextran [218]. Stimulation of hematopoiesis probably follows iron repletion. When hematopoiesis is derived primarily from PIG-A mutant stem cells, GPI-AP deficient erythrocytes dominate among newly synthesized cells, and the peripheral blood will be composed of a larger portion of complement-sensitive cells that will subsequently undergo hemolysis as a cohort. If a patient does experience clinically significant hemolysis in association with iron replacement, hematopoiesis can be suppressed by a brief period of transfusion or a short course of high-dose prednisone can be given during the early course of the iron replacement program to attenuate hemolysis. Splenectomy should be undertaken with caution because of the thrombophilia associated with PNH. Any operative procedure, including splenectomy, is associated with an increased risk of thromboembolic disease. Further, hemolysis is usually unaffected by splenectomy, although occasional responses have been reported (including one patient extensively studied by Ham [4, 9]). Patients who may be most likely to benefit from splenectomy are those with splenomegally, clinically significant cytopenias, and a cellular marrow. Such treatment may ameliorate the cytopenias primarily by eliminating sequestration. Therapy aimed either at inhibiting complement activation or at restoring the complement-regulatory proteins that are deficient on PNH erythrocytes would provide a more specific approach to management of the hemolysis of PNH. A recombinant, soluble form of complement receptor type I (sCR1) is currently in clinical trials for treatment of a variety of complement-mediated diseases.

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This protein is a potent inhibitor of both the classical and the alternative pathways [219], and a modified form that inhibits only the alternative pathway has been developed. Conceivably, sCR1 could be used to treat acute hemolytic episodes in patients with PNH. Its usefulness in management of the chronic complementmediated hemolysis of PNH, however, would be limited both by the current high cost and the requirement for intravenous infusion. Further, chronic suppression of the alternative pathway may result in a high incidence of infectious complications. The functional properties of recombinant, soluble MIRL (sCD59) have also being investigated [220]. However, sCD59 is unlikely to be an effective inhibitor of complement-mediated lysis in vivo; in the presence of plasma, the functional activity of the protein is lost as a consequence of binding to lipoproteins [220]. A DAF–MIRL chimera has been generated by recombinant technology, and in vitro studies have shown that the hybrid molecule retains inhibitory activity for both the C3/C5 convertase and the MAC [221]. If this bifunctional protein were active in vivo. it might have efficacy in the treatment of acute exacerbations of hemolysis in PNH. Rate of clearance, requirement for infusion, extent of immunosuppression, and cost will impact upon its usefulness in the treatment of the chronic hemolysis of PNH. Combining a complement-regulatory protein with specific antibody might enhance the therapeutic index by targeting the inhibitor to a specific tissue (e.g., combining MIRL with antiglycophorin would target RBC). The properties of a recombinant MIRL–antibody hybrid are being studied [222]. If the hybrid molecule functions like isolated MIRL, as few as 500 molecules/RBC may be effective in reducing hemolysis [105]. When the inhibitory molecule is bound to the membrane as part of an antigen–antibody complex, however, its functional properties may be altered. Depletion of plasma complement components using humanized monoclonal antibodies is another strategy for controlling complement-mediated injury. In this case, targeting components that are present in relatively low concentrations (e.g., constituents of the MAC) are more likely to produce an effect than aiming at proteins such as C3 and C4, which are present in the plasma in relatively high concentrations. Phase I and phase II studies using a recombinant, humanized monoclonal anti-C5 to ameliorate complement-mediated injury during cardiopulmonary bypass have shown efficacy [223, 224]. Such a reagent should attenuate hemolysis in PNH, but with limitations similar to those of the other recombinant protein inhibitors of complement discussed above. Another attractive strategy to inhibit complement activation in vivo is the development of small synthetic peptides that block the activity of the C3/C5 convertase or the MAC. Compstatin, a C3 binding peptide that was identified using phage display technology, is a potent inhibitor of complement activation [225], restricting convertase formation by interfering with the enzymatic activation of C3. Compstatin is scheduled to undergo testing in humans in the near future. Whether Compstatin is advantageous relative to sCR1 or humanized antiC5 will depend upon pharmacokinetics, toxicity profile, and cost. Hypothetically, peptides that mimic MIRL function could be identified and used to inhibit complement-mediated lysis by blocking MAC formation. The mutant PIG-A gene that causes the deficiency of the erythrocyte complement regulatory proteins DAF and MIRL in PNH has been identified (reviewed

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in [226]). Correction of the fundamental abnormality by using techniques of gene therapy appears feasible. In many cases, however, this approach may not be necessary (or desirable). Because the bone marrow and peripheral blood of essentially all patients is a mosaic of normal and PIG-A mutant cells, there exists the potential to use pluripotent stem cells derived from the patient to repopulate the marrow. Both DAF and MIRL are expressed on CD34⫹ hematopoietic stem cells. Using fluorescence-activated cell sorting, the phenotypically normal cells can be separated from stem cells with the PNH phenotype [227–229]. In theory, if an adequate quantity of CD34⫹, GPI-AP⫹ cells could be obtained and expanded by this method, the selected stem cells could be used to repopulate the marrow of the patient. Studies using syngeneic marrow transplant for treatment of PNH (discussed above) indicate that a marrow ablative-conditioning regimen will be required if this type of autologous transplant is to be successful [209]. A potential problem with autologous transplant is that phenotypically normal stem cells may be functionally abnormal. Both the primary and secondary clonogenic potential of hematopoietic progenitors of both CD34⫹, GPI-AP⫹ and CD34⫹, GPI-AP⫺phenotypes derived from the same patient are impaired to the same extent [230]. Thus, rescue from marrow-ablative therapy using PNH stem cells that are phenotypically normal may be unsuccessful. These findings are also relevant to gene therapy. Correction of the PIG-A mutation may restore expression of GPIAP, but not complement the defect that accounts for the abnormal clonogenic properties of the mutant stem cells population. A better understanding of both the nature of the bone marrow injury that predisposes to PNH and the basis of the selective pressure that results in the clonal expansion of PIG-A mutant stem cells is needed in order to design therapy aimed at treating the underlying disease process rather than at treating disease symptoms.

Summary and Conclusions The cause of hemolysis in PNH has been clearly defined. The observations of Stru¨bing, Hijmans van den Berg, and Ham laid the foundation for subsequent investigations that culminated in the identification of the complement-regulatory proteins, DAF and MIRL, that are deficient in PNH (Table 3-4). Characterization of those proteins has been important not only in illuminating the pathophysiology of PNH but also in defining the mechanisms by which normal human erythrocytes are protected against complement-mediated injury. As these widely expressed proteins are also involved in regulating complement activity in virtually all human tissues, both normal and malignant, and homologs have been identified in a variety of mammals, as well as in parasites and other microorganisms [56, 106, 231–236], pathophysiological studies of hemolysis in PNH have had a remarkably broad impact upon medicine and biology. Thoughtful interpretation of results from investigations into the abnormal lytic sensitivity of PNH erythrocytes has led to an understanding of most of the fundamental characteristic of the disease. The observation that not all erythrocytes from patients with PNH were susceptible to lysis in acidified serum suggested to Dacie [20] that PNH arose as a result of a somatic mutation. This

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TABLE 3-4 Observations That Helped Define the Basis of the Hemolysis of PNH Date

Observation

Ref.

1882

Nocturnal hemoglobinuria established as the defining clinical characteristics of the disease Hypothesis that hemolysis caused by accumulation of CO2 during sleep proposed Plasma observed to be red in color after severe attacks suggesting intravascular hemolysis Urine pigment identified as hemoglobin Plasma is normal Red cells are defective Lysis enhanced by acidification Hemolysis of PNH RBC in acidified serum mediated by a substance indistinguishable from complement Antibody not required for lysis of PNH RBC in acidified serum Discovery of the alternative pathway of complement Acidified serum lysis of PNH RBC mediated by the alternative pathway Complement-sensitive and -insensitive populations identified and characterized Intermediately sensitive (PNH II) phenotype identified Greater binding of C3 to PNH RBC observed Greater lysis of PNH RBC per C3 molecule bound Greater efficiency of the MAC on PNH RBC PNH III but not PNH II susceptible to reactive lysis Deficiency of DAF (CD55) reported Greater binding of C9 to PNH III cells in a reactive lysis system Greater lysis of PNH II and III RBC per C9 molecule bound Deficiency of MIRL (CD59) reported Basis of PNH II and PNH III phenotypes determined Basis of susceptibility of PNH RBC to acidified serum lysis characterized

[3]

1911

1939

1954 1955 1966 1973 1973 1978 1979 1983 1985 1989 1989 1991

[5]

[8]

[7] [19] [11] [134] [23] [140] [141] [154, 155] [96] [105] [138] [123]

concept, so elegantly articulated by Dacie 36 years ago, continues to guide current investigations into the origins of PNH. Careful analysis of the complement sensitivity of the erythrocytes by Rosse and colleagues [11, 134–137] confirmed that the peripheral blood of patients with PNH is a mosaic consisting of phenotypically normal and abnormal cells. Even more remarkably, more than one abnormal phenotype could be identified in many patients [134–137]. This unique characteristic became a defining feature of the disease and formed the basis for the concept that PNH does not arise simply by monoclonal expansion [134]. When techniques that were used to quantitate complement sensitivity of PNH red cells were applied to granulocytes and platelets [132, 133], the PNH defect was inferred to arise in a primitive hematopoietic stem cell that can differentiate along myeloid and megakaryocytic as well as erythroid lines. Thus, PNH was among the first diseases to be considered a disorder of the hematopoietic stem cell. The concept of a pluripotent hematopoietic stem cell was an issue of active debate in 1969 [132], and these findings of Aster and Enright supported

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the existence of a bone marrow stem cell. (It was not until 1976 that polycythemia vera was shown to be a stem cell disorder [237]). Although PNH erythrocytes were shown to be deficient in acetylcholinesterase in 1959 [127], it was the studies of Kunstiling and Rose [129] 10 years later demonstrating that the deficiency is confined to the complement-sensitive population that confirmed a pathological link with the disease. When DAF was found to be deficient, the absence of acetylcholinesterase took on even more importance because it suggested that the defect in PNH affected the expression of more than one protein. Earlier, PNH neutrophils were shown to be deficient in alkaline phosphatase [60]. When, in 1986, DAF was shown to be a member of the newly defined family of GPI-AP that included erythrocyte acetylcholinesterase and leukocyte alkaline phosphatase [130], the basis of the deficiency of multiple proteins in PNH became apparent. The observation that all proteins that are deficient in PNH are GPI-anchored led directly to the identification of PIG-A as the gene that is mutant in PNH [226]. Once the complement-regulatory proteins that are deficient in PNH were identified, the basis of the phenotypic mosaicism that was defined initially by using the complement lysis sensitivity assay became obvious [139]. By analyzing PIG-A mutations, phenotypic mosaicism was shown to be a consequence of genotypic mosaicism [139]. The existence of multiple mutant clones in the same patient [138] suggested that a powerful selective pressure was exerted on the bone marrow with the conditional growth or survival advantage being based on deficiency of one or more GPI-AP [238]. These observations support the basic tenets of the hypothesis that Dacie proposed in 1963 [20]. This summary highlights the important role that investigations of the hemolysis of PNH have played in defining the molecular basis of the disease. Are there other insights into the underlying pathophysiology that can be gained by studying the hemolytic component of the disease? It seems unlikely that additional GPIanchored erythrocyte complement-regulatory proteins will be identified as deficiency of DAF and MIRL appears to account entirely for the complement sensitivity of PNH RBC [123]. Conceivably, hemolysis may contribute to the thrombophilia of PNH [239]. Although there is no obvious correlation between the incidence of thromboembolic events and the proportion of GPI-AP deficient erythrocytes, there is a clinical impression that thrombosis occurs more often during hemolytic exacerbations [110], and this relationship warrants further investigation. Is the hemolysis of PNH strictly an epiphenomenon or does the deficiency of complement regulatory proteins play a role in the clonal dominance of the PIG-A mutant stem cells? In vitro studies suggest that sublytic amounts of complement can induce cellular proliferation [240, 241]. Whether this process is mediated through signaling by GPI-anchored complement-regulatory proteins [56, 106, 242, 243] has not been determined, and, it is unclear whether deficiency of DAF and MIRL enhances or obviates this process. Signalling by GPI-AP appears to be an important physiological process [244, 245], and its relationship to the clonal dominance in PNH needs to be clarified. In a 1963 review of PNH, Dacie [20] listed the following five central problems connected with PNH: (1) The nature of the red-cell defect. (2) The nature of the factors in normal plasma which bring about hemolysis of the PNH red

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cell. (3) Whether the patient’s leukocytes and platelets are abnormal. (4) The relationship between PNH and thrombosis. (5) The ultimate problem—the etiology of the disease and its relationship to marrow failure. The first three problems (and a number of others) have been solved, and in the process, a remarkable store of basic information has accrued. The last two questions remain challenges for the future. In addition to solving them, we must develop better treatment for the underlying process and for its complications.

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214. Rosse, W. F. (1989). Transfusion in paroxysmal nocturnal hemoglobinuria: To wash or not to wash. Transfusion 29, 663–664. 215. Hartmann, R. C., and Auditore, J. V. (1959). Paroxysmal nocturnal hemoglobinuria. I. Clinical studies. Am. J. Med. 27, 389. 216. Hartmann, R. C., Jenkins, D. E., Jr., McKee, L. C., and Heyssel, R. M. (1966). Paroxysmal nocturnal hemoglobinuria: Clinical and laboratory studies relating to iron metabolism and therapy with androgen and iron. Medicine 45, 331–363. 217. Rosse, W. F. (1997). Paroxysmal nocturnal hemoglobinuria as a molecular disease. Medicine 76, 63–93. 218. Parker, C. J., and Lee, G. R. (1998). Paroxysmal nocturnal hemoglobinuria. In ‘‘Wintrobe’s Clinical Hematology’’ (G. R. Lee, J. Foerster, J. Lukens, F. Paraskevas, J. P. Greer, and G. M. Rodgers, Eds.), 10th ed., pp. 1264–1288. Williams & Wilkins, Philadelphia. 219. Weisman, H. F., Bartow, T., Leppo, M. K., Marsh, H. C., Jr., Carson, G. R., Concino, M. F., Boyle, M. P., Roux, K. H., Weisfeldt, M. L., and Fearon, D. T. (1990). Soluble human complement receptor type 1: In vivo inhibitor of complement suppressing post-ischemic myocardial inflammation and necrosis. Science 249, 146–151. 220. Sugita, Y., Ito, K., Shiozuka, K., Suzuki, H., Gushima, H., Tomita, M., and Masuho, Y. (1994). Recombinant soluble CD59 inhibits reactive haemolysis with complement. Immunology 82, 34–41. 221. Fodor, W. L., Rollins, S. A., Guilmette, E. R., Setter, E., and Squinto, S. P. (1995). A novel bifunctional chimeric complement inhibitor that regulates C3 convertase and formation of the membrane attchk complex. J. Immunol. 155, 4135–4138. 222. Zhang, H., Yu, J., Bajwa, E., Morrison, S. L., and Tomlinson, S. (1999). Targeting of functional antibody-CD59 fusion proteins to a cell surface. J. Clin. Invest. 103, 55–61. 223. Thomas, T. C., Rollins, S. A., Rother, R. P., Giannoni, M. S., Hartman, Sl. L., Elliot, S. A., Nye, S. H., Matis, L. A., Squinto, S. P., and Evans, M. J. (1996). Inhibition of complement activity by humanized anti-C5 antibody and single-chain Fv. Mol. Immunol. 33, 1389–1401. 224. Rollins, S. A., Fitch, J. C. K., Shernan, S., Rinder, C. S., Rinder, H. M., Smith, B. R., Collard, C. D., Stahl, G. L., Alford, B. L., Li, L., and Matis, L. A. (1998). Anti-C5 single chain antibody therapy blocks complement and leukocyte activation and reduces myocardial tissue damage in CPB patients. Mol. Immunol. 35, 397 (abstr). 225. Sahu, A., Kay, B. K., and Lambris, J. D. (1996). Inhibition of human complement by a C3binding peptide isolated from a phage-display random peptide library. J. Immunol. 157, 884–891. 226. Kinoshita, T., Inoue, N., and Takeda, J. (1995). Defective glycosyl phosphatidylinositol anchor synthesis and paroxysmal nocturnal hemoglobinuria. Adv. Immunol. 60, 57–102. 227. Terstappen, L. W. M. M., Nguyen, M., Huang, S., Lazarus, H. M., and Medof, M. E. (1993). Defective and normal haematopoietic stem cells in paroxysmal nocturnal haemoglobinuria. Br. J. Haematol. 84, 504–514. 228. Prince, G. M., Nguyen, M., Lazarus, H. M., Brodsky, R. A., Terstappen, L. W., and Medof, M. E. (1995). Peripheral blood harvest of unaffected CD34⫹ CD38⫺ hematopoietic precursoners in paroxysmal nocturnal hemoglobinuria. Blood 86, 3381–3386. 229. Johnson, R. J., Rawstron, A. C., Richards, S., Morgan, G. J., Norfolk, D. R., O’Connor, S., and Hillmen, P. (1998). Circulating primitive stem cells in paroxysmal nocturnal hemoglobinuria (PNH) are predominantly normal in phenotype but granulocyte colony-stimulating factor treatment mobilizes mainly PNH stem cells. Blood 91, 4504–4508. 230. Maciejewski, J., Sloand, E., Sato, T., Anderson, S., and Young, N. S. (1997). Impaired hematopoiesis in paroxysmal nocturnal hemoglobinuria/aplastic anemia is not associated with a selective defect in the glycosyl phosphatidylinositol-anchored protein deficient clone. Blood 89, 1173– 1181. 231. Fishelson, Z. (1994). Complement-related proteins in pathogenic organisms. Springer Semin. Immunopathol. 15, 345–368. 232. Rother, R. P., Rollins, S. A., Fodor, W. L., Albrecht, J. C., Setter, E., Fleckenstein, B., and Squinto, S. P. (1994). Inhibition of complement-mediated cytolysis by the terminal complement inhibitor of herpesvirus saimiri. J. Virol. 68, 730–737. 233. van den Berg, C. W., and Morgan, B. P. (1994). Complement-inhibiting activities of human CD59 and analogues from rat, sheep, and pig are not homologously restricted. J. Immunol. 152, 4095–4101.

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234. Maio, M., Brasoveanu, L. I., Coral, S., Sigalotti, L., Lamaj, E., Gasparollo, A., Visintin, A., Altomonte, M., and Fonsatti, E. (1998). Structure, distribution, and functional role of protectin (CD59) in complement-susceptibility and in immunotherapy of human malignancies. Int. J. Oncol. 13, 305–318. 235. Saifuddin, M., Parker, C. J., Peeples, M. E., Gorny, M. K., Zolla-Pazner, S., Ghassemi, M., Rooney, I. A., Atkinson, J. P., and Spear, G. T. (1995). Role of virion-associated glycosylphosphatidylinositol-linked proteins CD55 and CD59 in complement resistance of cell line-derived and primary isolates of HIV-1. J. Exp. Med. 182, 501–509. 236. Spear, G. T., Lurain, N. S., Parker, C. J., Ghassemi, G., Payne, G. H., and Saifuddin, M. (1995). Host cell-derived complement control proteins CD55 and CD59 are incorporated into the virions of two unrelated enveloped viruses: Human T cell leukemia/lymphoma virus type I (HTLV-I) and human cytomegalovirus (HCMV). J. Immunol. 155, 4376–4381. 237. Adamson, J. W., Fialkow, P. J., and Murphy, S., (1976). Polycythemia stem-cell and probable clonal origin of the disease. N. Engl. J. Med. 295, 913–916. 238. Bessler, M., Mason, P., Hillmen, P., and Luzzatto, L. (1994). Somatic mutations and cellular selection in paroxysmal nocturnal haemoglobinuria. Lancet 343, 951–953. 239. Ninomya, H., Kawashima, Y., Hasegawa, Y., and Nagasawa, T. (1998). Microvesicles generated from PNH red blood cells by complement activation contribute to the thrombotic diathesis in PNH. Blood 92 (supplement), 155a (abstr). 240. Niculescu, R., Bades, T., and Rus, H. (1998). Sublytic C5b-9 induces proliferation of human aortic smooth muscle cells. Mol. Immunol. 35, 400 (abstr). 241. Rus, H., Soane, L., Niculescu, F., and Shin, M. L. (1998). Oligodendrocyte protection by sublytic C5b-9 from apoptotic cell death. Mol. Immunol. 35, 400 (abstr). 242. Korty, P. E., Brando, C., and Shevach, E. M. (1991). CD59 functions as a signal-transducing molecule for human T cell activation. J. Immunol. 146, 4092–4098. 243. Murray, E. W., and Robbins, S. M. (1998). Antibody cross-linking of the glycosylphosphatidylinositol-linked protein CD59 on hematopoietic cells induces signaling pathways resembling activation by complement. J. Biol. Chem. 273, 25279–25284. 244. Lublin, D. M. (1992). Glycosyl-phosphatidylinositol anchoring of membrane proteins. Curr. Top. Microbiol. Immunol. 178, 141–162. 245. Robinson, P. J. (1991). Phosphatidylinostiol membrane anchors and T-cell activation. Immunol. Today 12, 35–41.

4 Thrombotic Complications in PNH Elaine M. Sloand and Neal S. Young National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892-2490

Introduction Paroxysmal nocturnal hemoglobinuria (PNH), a clonal stem cell disorder, is associated with an increased risk of venous thrombosis. Thrombosis is responsible for significant morbidity and is the complication most clearly associated with poor prognosis, including death, in clinical studies [1, 2]. The pathophysiology of thrombosis in PNH is not understood. No coagulation or platelet abnormality has been identified that can adequately account for the hypercoagulable state observed. Moreover, good case-controlled epidemiologic studies of this rare disorder are lacking, and those limited studies that are available show unexplained geographic variation in the incidence of thrombosis and a preponderance of abdominal venous thrombosis not seen in other cases of thrombophilia. A general impediment in our ability to comprehend thrombosis in PNH is the failure to account for and understand the pathophysiology of most thrombophilia quite independent of this disease association. In 60% of all venous thrombosis, no cause or risk factor can be identified. Furthermore, PNH, a disorder associated with cellular abnormalities, is distinctly different from the other conditions associated with venous thrombosis. With the exception of the antiphospholipid syndrome, all other syndromes with increased risk of venous thrombosis are related to defects in clotting proteins and factors. Despite intensive investigation, no defect or abnormality in coagulation has been identified in PNH [3]. Compounding difficulties, the PNH patient populations studied are heterogeneous: patients may present with aplastic anemia or with classical hemolytic PNH, some profoundly symptomatic, others with only subtle flow cytometry abnormalities. The results of both clinical and basic studies are difficult to interPNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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pret when the population under study has not been well described or characterized. This chapter will review the limited data available on PNH-related thrombosis, and the defects in glycosylphosphatidylinositol (GPI)-linked proteins as they related to coagulation, and formulate the important questions that remain regarding thrombotic complications of this rare disease.

Incidence of Thrombosis in PNH The incidence of thrombosis among patients with PNH varies greatly. While thrombosis is a frequent complication of PNH among European cohorts, it is relatively rare in Asian populations. A retrospective study of a group of 80 PNH patients in the United Kingdom [2] demonstrated that 39% experienced at least one episode of thrombosis during the course of their disease and many experienced several events; thrombosis was the cause of death in 30% of cases. Sites of thrombosis included peripheral as well as intraabdominal veins (Table 4-1) and Budd-Chiari syndrome was relatively common. Arterial thrombosis in this group of patients was rare and the rate may not be elevated over that seen in the general population. The presence of thrombosis was similar in a large multicenter French study [4], and in a series of patients collected in Spain [5] and Mexico [6]. In marked contrast, thrombosis was unusual in Japanese [1], Chinese [7], and Thai [8] populations. As can be seen in (Table 4-2) [2], the occurrence of thrombosis appears to be most closely related to country of origin, although some cohorts with a greater proportion of patients presenting with aplastic anemia also had inversely fewer thrombotic episodes. The lower frequency in Asians may be explained in part by the globally much lower rate of thrombotic disease in

TABLE 4-1 Sites and Types of Thrombosis a No. of patients Intraabdominal Hepatic vein Inferior vena cava Mesenteric vein Splenic vein Renal vein Unspecified Other venous sites Cerebral vein Pulmonary embolism Deep vein Superficial Arterial Myocardial infarction Cerebrovascular accident

8 3 4 1 1 1 4 9 7 3 6 2

Adapted from [2], with permission. Copyright © 1995 Massachusetts Medical Society. All rights reserved.

a

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TABLE 4-2 Frequency of Thrombosis in Patients with PNH and Relationship to Numbers Presenting with Aplastic Anemia and Country of Origin Investigator

Location

% thrombosis

% presenting with AA

Hillmen [2] Dunn [44] Forman [45] Gongora-Biachi [6] Le [46] Fujioka [1] Krauatrachue [8] Socie [4] Young

United Kingdom Taiwan United Kingdom Mexico China Japan Thailand France United States

39 7 31 23 2 2 2 27 6

23 17.5 16 45 38 41 40 30 85

these ethnic groups, which may be related in turn to diet, environmental factors, and especially genetic factors. In one study of the general U.S. population, Asian and Pacific Islanders had 1/4th the incidence of venous thrombosis compared to Caucasians and African Americans [9]. There are also important differences in the nature of the PNH patient populations studied. In Asian clinics there may be selection for patients with a preexisting history of aplastic anemia (as also in our referral center in Bethesda). Whether there are differences in the PIG-A mutations in European cohorts that predispose to thrombosis is speculative, but to date, unsupported by laboratory evidence. Finally, the effect of treatment, including antithymocyte globulin (ATG) or cyclosporin, on the incidence of thrombosis has also not been investigated.

Factors Associated with Venous Thrombosis Risk factors for venous thrombosis in individuals without PNH have not been well defined despite substantial research, and as stated above, in the majority of cases no determinants can be identified. Activated protein C resistance due to the Leiden mutation, ATIII deficiency, protein C deficiency, protein S deficiency, homocysteinuria, prothrombin gene mutation, and inherited abnormalities of fibrinolysis are all hereditary factors associated with venous thrombosis (Table 4-3). ATIII, protein C, and protein S deficiencies can be found in fewer that 5% of patients with thromboembolism, while factor V Leiden and the prothrombin gene mutations are more common and present in a large proportion of patients with no other predisposing factors [10, 11]. A number of acquired conditions including cancer, recent surgery, and oral contraceptive use are also associated with venous thrombosis. An additional acquired disorder, the antiphospholipid antibody syndrome, has been linked to as many as 15–20% of all venous thromboembolic events [12]; the antibody is believed to interfere with activation of factor Va, leading to resistance to activated protein C. While defects or abnormalities of coagulation predispose to venous thrombosis, to date, platelet or leukocyte dysfunction has not been implicated in predisposing to clot formation in veins. PNH, which results from the absence of the GPI anchor and leads to well-defined cellular membrane defects, is clearly an exception to all other conditions traditionally associated with venous thrombosis.

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TABLE 4-3 Risk Factors Associated with Arterial and Venous Thrombosis Venous Factor V Leiden ATIII deficiency Protein C deficiency Protein S deficiency Prothrombin G20210A gene mutation Dysfibrinogenemia Hyperhomocysteinemia Pregnancy, estrogen use Immobilization, trauma, surgery Antiphospholipid syndrome Malignancy, nephrotic syndrome, myeloproliferative syndrome

Arterial Smoking Hypertension Obesity Diabetes Reduced fibrinolytic function Hyperhomocysteinemia Inflammatory markers (fibrinogen, C-reactive protein, serum amyloid A) Hyperlipidemia Platelet aggregation, activity, function

Platelet Abnormalities in PNH Platelets of patients with PNH do demonstrate abnormalities in GPI-linked proteins. GPI-linked proteins present on platelets are CD55, CD59 [13], urokinase plasminogen activator receptor (uPAR) [14], GP500, and GP175 [15]. Except for uPAR, it is not clear that any of these proteins has any functional significance with respect to coagulation. In a study of 10 PNH patients conducted in our laboratory, [13] abnormalities of CD55 and CD59 were found in all cases (Fig. 4-1, Table 4-4).

Figure 4-1 Expression of CD55 and CD59 on platelets in PNH. Normal platelets (A and B) and those derived from patients with PNH (C and D) were stained with CD42b and CD59 (A and C) and CD42b and CD55 (B and D). Panels A–D represent two-color diagrams with log red fluorescence intensity on the ordinate and green fluorescence intensity on the abscissa. PE, phycoerythrin; FITC, fluorescein isothiocyanate.

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TABLE 4-4 Expression of GPI-Linked Molecules on Platelets a

PNH1 PNH2 PNH3 PNH4 PNH5 PNH6 PNH7 PNH8 PNH9 PNH10 AA1 AA2 AA3 AA4 AA5 AA6 AA7 N ⫽ 20

Monocytes

Neutrophils

Ham test

CD 14

CD 16

CD 55

CD 59

20% 61% (-) 34% 11% 20% 4% 40% 2% 31%

4 1 39 18 22 5 26 0.8 40 11

44 62 71 33 22 10 86 1.2 71 70

63 49 83 59 32 62 55 30 54 53

44 60 91 56 n.d. b 10 45 n.d. b 11 88

93 83 83 80 92 87 86

90 94 93 96 97 87 92

88 92 86 88 80 90 90

90 95 83 80 60 70 70

85 ⫾ 6

93 ⫾ 3

(-) (-) (-) (-) (-) (-) (-) 1(-)

83 ⫾ 6

90 ⫾ 4

Platelets

a

Cells were stained with antibodies to GPI-linked portions and to lineage-specific antigens. For monocytes, staining was performed with CD13 antibody and CD14 antibody in the monocytic gate and expressed as percentage of CD14⫹ cells with CD13⫹ cells. For neutrophils, staining was performed with CD14 and CD15 antibodies and expressed as percentage of CD16⫹ within CD15⫹ cells in the granulocyte gate. For platelets, staining was performed using CD55 and CD59 antibodies and expressed as percentage of CD55/CD59⫹ within CD42b⫹ platelets. In PNH patients an average of 17 ⫾ 5% of monocytes expressed CD14 antigens, 40 ⫾ 10% of neutrophils expressed CD16 antigens, and 54 ⫾ 5% of platelets expressed CD55 antigen. b n.d., not done.

Expression of activation markers, CD62 and CD63, in unstimulated PNH platelets were comparable to normals, as was the cell surface content of two other membrane-associated proteins, GPIb and GPIIb/IIIa. uPAR, while present on activated normal platelets, was absent on platelets of the PNH phenotype (Fig. 4-2). Consistent with these results, CD55, CD59, and uPAR could be removed from normal platelets by treatment with phospholipase C (PIPLC). However, platelets from PNH patients did not undergo spontaneous aggregation, nor were they more sensitive to aggregating stimuli. Platelets with and without the PNH phenotype expressed activation markers equally well after stimulation with a number of agonists, including, ADP, collagen, and epinephrine [13]. Activation did not affect the expression of CD55 or CD59. One group of investigators examined the effects of complement on platelet release of serotonin [16]. When complement was activated via the alternative pathway, more C3 was fixed to the PNH platelet than to the normal platelet. While the PNH platelet degranulated after activation by C3 fixation, normal

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Figure 4-2 Expression of uPAR on unactivated and activated normal and PNH platelets. Platelets from normals and patients with PNH were activated with ADP and with epinephrine and stained with CD41-TC and uPAR-FITC. Cells were gated to include only platelets staining with CD41-TC. Cells staining (phycoerythrin tricolor) with uPAR-FITC are seen above. Eighty percent of the PNH patient’s platelets lacked DAF.

platelets required fixation of the terminal components of complement. Maximal release of serotonin was achieved in the presence of the terminal elements of complement in PNH platelets, but this was probably the result of membrane damage. The sensitivity of PNH platelets to complement as well as the increased tendency of PNH platelets to form microvesicles (sealed membrane fragments with procoagulant activity) [17] are of possible clinical importance. In clinical correlation, thrombotic episodes have been described in association with periods of hemolysis, which presumably reflect complement activation. Although Aster and colleagues showed that the PNH platelet, like the PNH red cell, was more sensitive to lysis by complement in vitro than normal platelets, other investigators have demonstrated normal platelet survivals in individuals


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with PNH [18]. Normal 111Indium-labeled platelet survivals may be explained by the fact that, unlike other blood cells, platelets have additional mechanisms for regulating the alternative pathway of complement. For example, platelets release factor H, a protein that regulates complement, in response to C3 fixation. Factor H is found in the platelet alpha granule and appears in response to both C3 fixation and thrombin stimulation. In a study of 19 patients, platelets from PNH patients had normal C3 convertase activity, despite the absence of DAF on 80–95% of the cells [18]. PNH patient platelets released less factor H than did normal platelets, and the amount was lowest in patients with elevated C3 convertase activity, implicating a role for factor H in complement regulation. Other complement-regulatory proteins that have been described in platelets include C1 inhibitor [19] and factor D [20].

Role of Red Cell and Platelet Microvesicles Platelet and red cell fragments form microvesicles with procoagulant activity, but their role in coagulation and thrombosis has been disputed [21]. Platelet microvesicles arise after activation and exposure to complement; they bind activated factor V and factor VIII as well as protein C inhibitor [22]. Microvesicles can support hemostasis in vitro and inhibit the action of protein C. When platelet microvesicles were infused into thrombocytopenic rabbits they acted as a platelet substitute [23]. Microvesicles are increased in a number of clinical conditions accompanied by platelet activation, as in thrombotic thrombocytopenic purpura (TTP) [24] and after cardiac bypass [14]. However, too few microvesicles circulate in the plasma of normal persons to play a systemic role in coagulation, although they might be important locally at areas of endothelial injury. One case of a bleeding diathesis was attributed to a congenital defect in the ability of platelets to form vesicles [25]. In our study of patients with PNH, the numbers of microvesicles before and after activation with the agonists ADP and collagen were similar to those of controls [13]. In addition, the microvesicles formed bound normal amounts of factor V and factor VIII. However, these results cannot be extrapolated to all PNH, as our patients mainly presented with aplastic anemia and only one had a history of thrombosis. Both PNH red cells and platelets appeared defective in their ability to form vesicles in response to the calcium ionophore A23187 compared to a normal response to platelet agonists [26]. Mutant Blymphoyblastoid cell lines, lacking GPI-linked proteins, produced about half the number of microvesicles compared to wild type. Possibly the ability to form vesicles helps the cell eliminate portions of the membrane which are bound to activated complement, conferring a protective advantage. However, the PNH platelet may behave differently from the normal platelet when exposed to complement. Excessive numbers of procoagulant-rich microvesicles were formed by platelets with the PNH phenotype after exposure to complement [17]. These findings, along with the previously discussed sensitivity for the complement-induced release reaction, are of interest because thrombotic episodes have been described to accompany episodes of hemolysis.

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Urokinase Activator Receptor (uPAR) and Cell-Mediated Fibrinolysis The role of the urokinase activator receptor (uPAR), a GPI-linked protein, in cell-mediated fibrinolysis has not been substantially investigated and its clinical significance remains in doubt [27, 28]. uPAR serves as a receptor for the inactive zymogen prourokinase activator (pro-uPA), which is converted to its active form uPA subsequent to binding to uPAR. Active uPA then converts inactive plasminogen to active plasmin (Fig. 4-3). uPA that is bound to uPAR generates 14-fold more fibrinolytic activity compared to uPA alone [29]. In addition, membrane-associated plasmin is protected from 움-1 anti-plasmin, which readily inactivates free plasmin. The strong acceleration of activation of plasminogen by uPAR is also related to feedback activation of pro-uPA by cell-bound plasmin (Fig. 43); uPAR, lowers the Km value of the uPA-mediated activation of plasminogen to a level below the physiologic concentration of plasminogen. Activated normal leukocytes and platelets express uPAR on their membrane surface. In vitro studies demonstrate that the fibrinolytic activity produced by uPA is increased in the presence of leukocytes [30]. Leukocytes with the PNH phenotype do not express uPAR on their surface, and these cells are defective in their ability to support fibrinolysis [30]. Similarly, unpublished results from our laboratory show that uPAR is expressed on activated platelets from normal individuals; other investigators demonstrated that thrombinstimulated normal platelets produced active plasmin from plasminogen [31]. We have found that uPAR was not expressed on platelets from patients with PNH, and this inability does not appear to be related to a defect in protein synthesis; uPAR mRNA is present in PNH leukocytes and appears to be normal [30]. In addition, patients with PNH have been reported to have greatly increased plasma levels of soluble uPAR [32, 33]. Soluble uPAR may

Figure 4-3 Mechanism of action of uPAR: relationship between binding of uPA and conversion of plasminogen to plasmin. Pro-uPAR is activated by trace amounts of plasmin (PL) after binding to uPAR. Active uPA will subsequently activate plasminogen (PLG) to generate more plasmin. Plasminogen activator inhibitors (PAI-1 and PAI-2) may directly inhibit uPA and downregulate the activity of the system. Reproduced with permission from Stem Cells 15, 398–408 (1977); copyright by AlphaMed Press.


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compete with cell-associated uPAR for binding to uPA. It may be hypothesized that uPAR is synthesized by the PNH cell, but is incapable of membrane attachment because of the absence of the GPI anchor. However, uPAR is present on vascular smooth muscle cells, unaffected by any mutation in hematopoietic stem cells and where its expression could be induced by thrombin [34]. The larger question is whether cell-mediated fibrinolysis is of clinical relevance. Certainly defects in noncell-associated fibrinolysis have been linked to thrombotic disease [35, 36]. However, the uPAR knock-out mouse did not demonstrate any abnormalities in clotting, although the double TPA/ uPAR knock-out mouse showed excessive fibrin deposits in the hepatic vein [37]. While knock-out animal models may not always recapitulate human pathology, this work seriously calls into question the importance of uPAR acting alone in the prevention of thrombosis.

Treatment of Thrombosis in PNH Definitive recommendations regarding the treatment of thrombosis in PNH cannot be stated, as no studies have clearly compared various regimens in this rare disease, and there is no clear pathophysiologic foundation for the thrombotic proclivity. Despite the large number of new drugs available that block coagulation, it is the opinion of experts that most cases of PNH venous thrombosis should still be treated with standard anticoagulation regimens. The number of patients who currently fail treatment with conventional anticoagulation is not known, but such failures are frequent in many published series [2, 4]. Heparin has even been shown to potentiate complement activation in vitro [38]. Despite some meager evidence that excessive platelet activation secondary to complement may be involved, no clinical study has been made of platelet inhibitors such as aspirin or GPIIb/IIIa blockers. If complement attachment on platelets were important in generating thrombosis, anti-CD5a mAb might prove therapeutically effective. Thrombolytic agents were reported to be helpful in four cases of established Budd-Chiari syndrome [39, 40]; these patients regained liver function and were asymptomatic after long-term follow-up. Liver transplantation has been successful in restoring normal liver function in one patient [41]. Bone marrow replacement has been undertaken in two identical twins who were reportedly free of disease on long-term follow-up [42, 43]. With the meager available clinical data on treatment and without a clear understanding of the pathogenesis of clot formation in PNH, it is difficult to develop coherent treatment strategies.

Summary While no defect in GPI-linked protein has been definitively shown to increase clot formation, platelet sensitivity to complement and the absence of cell-associated uPAR have been identified as candidate lesions. Analysis of knock-out animal models may lead to a better understanding of the factors leading to thrombosis.

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Better epidemiologic studies are needed to understand both the extent and the basis of the geographic and ethnic disparities in thrombosis rate. Most importantly, knowledge of the pathophysiology of this so frequently devastating condition should lead to rational approaches to treatment. Individuals who are at greatest risk for thrombosis could be identified early and availed of experimental or high-risk treatments.

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18.

19. 20.

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24. 25.

26. 27. 28. 29.

30.

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5 Bone Marrow Failure in PNH Daniel E. Dunn, Johnson M. Liu, and Neal S. Young Hematology Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892

Introduction Almost 100 years elapsed between the first descriptions by Dacie and Lewis [1, 2] of paroxysmal nocturnal hemoglobinuria PNH [3–5] and the clinical association of this hemolytic syndrome with bone marrow failure, specifically aplastic anemia. Initially, the diagnosis of PNH relied on the finding of intermittent ‘‘hemoglobinous urine (predominantly) in the morning [4].’’ With the advent of the Ham test [6, 7], it became possible to diagnose PNH in non-‘‘classical’’ cases in which hemoglobinuria might not be apparent; the case definition thus comprised a positive Ham test and clinical evidence of hemolysis, thrombosis, or bone marrow failure. The insensitivity of the Ham test, however, placed constraints on the diagnosis of PNH, especially in patients receiving red cell transfusions. In the modern era, flow cytometric techniques have proven highly sensitive and specific in the detection of the glycosylphosphatidylinositol (GPI)anchored protein (AP)-deficient leukocyte counterparts of PNH red cells. With the widespread application of these techniques to the analysis of patients with myriad hematopoietic disorders, it is reasonable to expect that a broader, more detailed picture of the spectrum of PNH will become apparent over the next few years. The molecular bases for the hemolytic and thrombotic manifestations in PNH have been traced to the GPI-AP-deficient phenotype (Fig. 5-1), and in particular the absence of decay-accelerating factor (DAF) (CD55) and membrane inhibitor of reactive lysis (MIRL) (CD59) on erythrocytes [8, 9], platelets [10–14], and possibly leukocytes [15] (see preceding chapters devoted to these issues). This phenotype is due to acquired mutations in an X-linked gene, PIG-A [16–19], PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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Figure 5-1 Structure of glycosylphosphatidylinositol-anchored proteins (GPI-AP). An oligosaccharide consisting of inositol (Ino), glucosamine (GlcN), and three mannoses (Mann1–3), plus a linking ethanolamine (EtN), bridge the terminal amino acid of a given protein (‘‘aa웆’’) by an ethanolamine (EtN) to the alkyl-acyl glycerol moiety which anchors the resultant GPI-AP in the cell membrane. In PNH, somatic mutations in the PIG-A gene (located on the X chromosome) result in defective GPI anchor biosynthesis. Cell surface expression of all proteins dependent on this mode of membrane anchoring is lost in the progeny of affected hematopoietic stem cells (HSC).

which is necessary for GPI-anchor biosynthesis [20]. These mutations occur in individual hematopoietic stem cells, the progeny of which eventually account for a large proportion of the hematopoietic activity in a patient at the time of presentation (see below). This chapter will focus on the large area of overlap between the syndromes of PNH and bone marrow failure and potential unifying pathophysiologic mechanisms suggested by the association of these two rare diseases. The central etiologic dilemma in PNH is why a PIG-A mutant hematopoietic stem cell (HSC) expands dramatically to account, in most cases, for the bulk of hematopoiesis, without inevitable progression to complete replacement of wildtype (GPI-AP-positive) hematopoiesis, polycythemia, or leukemia. The reciprocal question might just as easily be posed: depending on estimates of HSC number and the expected mutation rate, why does not everyone develop PNH? The key to answering these questions may rest in examining the causal relationship between PIG-A mutations and bone marrow failure, in particular, aplastic anemia

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(AA). Only about a third of patients with PNH will have received a formal diagnosis of AA prior to their physician’s recognition of PNH; another third, however, subsequent to their diagnosis with PNH, eventually fulfill the criteria for severe or moderate AA at some point over years to decades. Even the remaining ‘‘hemolytic’’ PNH patients have evidence of bone marrow failure in in vitro assays of hematopoiesis (see below). In the following sections, clinical, experimental, and theoretical observations on PNH patients, PIG-A mutations, and HSC biology will be presented, and a Darwinian model of PIG-A mutant HSC evolution in the context of selection by the processes underlying bone marrow aplasia will be proposed.

Pathophysiology of Marrow Failure in PNH Clinical Behavior of PNH and Studies with Patient Samples Stem Cell Clonality Inferences from observation of hematopoiesis in vivo and in vitro have been useful in providing insight into the basic pathologic processes at work in PNH. First, PNH is a clonal disease [21] and the ontogenic stage of the genetic lesion is at or before the pluripotent hematopoietic progenitor; in many PNH patients, every lymphoid and myeloid lineage harbors GPIAP(⫺) cells: T cells, B cells, natural killer (NK) cells, granulocytes, monocytes, platelets, and erythrocytes [22]. The advantage conferred by the PIG-A defective phenotype, whether intrinsic or extrinsic, must occur at the level of stem cell biology, although it is possible that selection may also be applied to more mature progenitors. (But progenitors, by definition, do not self-renew and therefore are not capable of making an enduring contribution to hematopoiesis.) Second, the natural history of PNH is neither inexorable expansion of the mutant clone, nor inevitable progression to leukemia. Rather, evolution to bone marrow failure is the second leading cause of death in PNH patients (after thrombosis) [23]. Moreover, in one longitudinal study among PNH patients who survived at least 10 years, 15% had a spontaneous remission [23]. Serial measurements of the percent of neutrophils deficient in GPI-AP expression in our patients have revealed very little change over almost 2 years of periodic cytofluorimetric monitoring (unpublished observations). Even when PNH patients develop chromosome abnormalities and are thus considered to have evolved into myelodysplasia (MDS), this cytogenetically distinctive clone does not inevitably coincide with the GPI-AP(⫺) population [24–28]. Results of Therapeutic Stem Cell Transplant Bone marrow transplantation appears to cure PNH—but only if preceded by a cytotoxic drug regimen. Simple infusion of syngeneic bone marrow is generally not curative [29–35] (Table 5-1). Nonablative transplants from identical twins have been attempted only when PNH has evolved to bone marrow failure, but clinical remissions, if achieved, are rarely permanent. In one recently reported case, a patient experienced a clinical remission for 10 years before symptoms recurred [30], and the PIG-A mutation that was identified at relapse was different from that found at diagnosis a decade earlier [30]. This case suggests that PIG-A mutant HSC may undergo clonal ‘‘exhaustion’’ (as likely do normal HSC), and expansion of a PIG-A

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TABLE 5-1 Summary of Outcomes of Syngeneic Marrow Transplants in PNH Fefer [33] Hershko [35] Jehn [34] Kolb [31] Kawahara [29, 36] a Endo [32] a

Recovery, late relapse PNH Recurrent PNH Persistent PNH Persistent PNH 1. Transient recurrent Ham(⫹); later thromboses with Ham(⫺) 2. Recurrent hemolysis, persistent Ham(⫹) Recurrent PNH

Patient #1 has been reported twice [29, 36].

mutant HSC clone to a clinically evident level may be a very slow process, perhaps requiring years. Furthermore, because the twin donor did not develop PNH or molecular evidence of the posttransplant PIG-A mutation, host pathology can be inferred as necessary for selection of PIG-A mutant HSC. The preparative regimen used in transplanting PNH patients is sometimes simply high-dose cyclophosphamide, and relapse after such transplants is rare [36, 37]. High-dose cyclophosphamide, without infusion of allogeneic HSC, appears to be therapeutic in AA[38] without stem cell rescue. This regimen has been applied in PNH; two PNH patients treated at the NIH, however, showed no improvement in the percentages of their GPI-AP(⫹) neutrophils after recovery of their counts (unpublished observations). Treatment of cases of PNH/AA with conventional ATG and/or CsA, while frequently improving blood counts, does not necessarily result in an increased proportion of normal [GPI-AP(⫹)] hematopoiesis (unpublished observations). At least two explanations may be offered to explain failure of the PIG-A mutant clonal contribution to regress. First, multiple inhibitory processes may suppress hematopoiesis in PNH/AA, only one of which depends upon GPI-AP expression by the target HSC; the process reversed by ATG and CsA would then necessarily be GPI-AP independent (an example of a latter such process might be IFN-웂- or TNF-움-mediated marrow suppression)[39, 40]. Alternatively, at the level of normal stem cell biology, in the absence of pathologic inhibition of hematopoiesis PIG-A mutations in the stem cells may be entirely neutral, and the relative levels of mutant and wild-type hematopoiesis prior to and after therapy would then be expected to remain unchanged. This second explanation, is consistent with the natural history of untreated PNH only if either the driving pathologic disease process frequently ‘‘burns out’’ at some point short of complete HSC replacement or if the intensity of the pathologic inhibition varies inversely with the level of hematopoiesis contributed by GPI-AP(⫹) HSC. While the period of time over which we have followed GPI-AP(⫺) proportions in PNH patients has been relatively short, retrospective cross-sectional data from our AA patients support these inferences, as patients with AA/PNH analyzed at longer follow-up intervals, up to 12 years, do not inevitably have higher fractions of PIG-A-mutant hematopoiesis than do patients studied much earlier in the course of their disease. In Vitro Colony Culture Studies A component of bone marrow failure is evident in most cases of PNH, either clinically, as manifested by bone marrow


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hypocellularity or by thrombocytopenia [41, 42], or experimentally, as reflected by in vitro colony assays performed on cells from even hypercellular bone marrow aspirates [43–47]. Individual patients with PNH represent a convenient source of hematopoietic progenitors that are chimeric with regard to the presence of PIG-A mutations but which have arisen from an identical genetic background and presumably comparable microenvironments. Fluorescent-activated cell sorting (FACS) has been utilized to separate wild-type GPI-AP(⫹) progenitors from PIG-A mutant GPI-AP(⫺) progenitors. When such progenitors have been assayed in various in vitro assays of hematopoiesis, such as colony formation [(by myeloid and erythroid progenitor cells (CFU-GM or BFU-E, respectively)] or the long-term culture-initiating cell assay (LTCIC), no significant difference in clonogenicity between these two subpopulations has been observed [48]. As mentioned above, however, the overall levels of colony formation, whether primary or secondary, are typically low in PNH [43–47] compared to normal controls. If expansion of PIG-A mutant HSC clone(s) in cases of PNH represents a selected adaptation of the bone marrow to external stress [49–51], then why is the growth of isolated GPI-AP(⫺) progenitors impaired? And, when the proportion of hematopoiesis contributed by PIG-A mutant clone(s) exceeds 80 or 90% (as is common in cases of hemolytic PNH), why is any diminution in progenitor numbers observed with unsorted bone marrow samples? Several conceptual and methodological considerations may account for the data. First, over a 2-week in vitro assay, the difference in growth between PIG-A mutant and wild-type progenitors may fall below the level of precision of typical colony count experiments (20–30%); a small difference compounded over a year or more, however, could ultimately result in dramatic expansion of a mutant clone. Second, the equivalent poor growth of GPI-AP-negative and positive progenitors in vitro may reflect a balance of two unrelated phenomena: PIG-A mutations may actually confer a modest disadvantage in vivo which may be exacerbated in vitro (for example, the folate receptor is GPI-anchored); the poor growth of GPIAP(⫹) progenitors in vitro, on the other hand, may represent the residual effect of their in vivo inhibition. Finally, it is possible that PIG-A mutations may indirectly result in aberrant or delayed expression of differentiation markers; in such a case, comparison of numbers of colonies formed by GPI-AP-positive or -negative CD34(⫹) cells, despite being derived from the same bone marrow sample, would not be appropriate. Human Hematopoiesis in Animal Models The relative growth of human GPI-AP(⫹) and GPI-AP(⫺) hematopoietic cells has also been examined in a surrogate in vivo assay system, the severely immunodeficient (SCID) mouse [52]. ‘‘Engraftment’’ of intravenously injected BM samples from PNH patients in these partially immunodeficient mice (SCID mice actually possess augmented NK activity) was variable from donor to donor, but whatever human cells did engraft they were found consistently to be GPI-AP(⫺). The authors inferred from these results that PIG-A mutant HSC from PNH patients possess an undefined intrinsic growth advantage, which they postulated was due either to the PIG-A mutation itself or additional mutation(s) acquired by the PIG-A mutant clone. The results with normal bone marrow samples in this study, however, differ from other published studies of human hematopoiesis in immunodeficient mice

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[53–55], as the investigators were unable to obtain engraftment of BM inocula from healthy subjects, and the human cells that did engraft from the PNH patients did not retain any CD34 expression. SCID/NOD mice, which are NK deficient, are required in order to obtain optimal engraftment of control normal adult bone marrow [53]. Thus, the failure of BM inocula from normal subjects to engraft in SCID mice could be due solely to active destruction by NK cells shortly after the intravenous injection and not to any intrinsic growth disadvantage of PIG-A normal progenitors. Were the interaction of mouse NK cells with human hematopoietic targets dependent on target GPI-AP expression, PNH BM would show a selective advantage in SCID mice. In the above-described growth experiments utilizing sorted GPI-AP(⫹) and GPI-AP(⫺) cells derived from patients, there is an implicit assumption that the only differences between the two populations relate to PIG-A gene function. Ideally, one would experimentally impose PIG-A mutant or wild-type status upon an otherwise homogeneous collection of primitive hematopoietic progenitors and then compare the behavior of such cells in assays of hematopoiesis. Paired Cell Lines The effects of experimentally manipulating PIG-A expression has been examined in cells other than hematopoietic progenitors, namely, Epstein-Barr virus (EBV)-transformed B lymphoblastoid cell lines (LCL) using cloned PIG-A mutant, GPI-AP(⫺) LCL from PNH patients [56, 57]. The first report [58] compared the relative sensitivities of the parental GPIAP(⫺) LCL with its PIG-A-transfected GPI-AP(⫹) daughter line with regard to various modes of programmed cell death. With two independent GPI-AP(⫺) LCL, reconstitution of PIG-A function appeared to result in restoration of susceptibility to apoptosis induced by either serum deprivation or 웂 irradiation. Transfection of the GPI-AP(⫺) LCL with a control vector was not described [58]. The second report of such experiments utilized two independent PIG-A expression vectors (one retroviral, the other EBV-based), as well as control vectors, and came to a different conclusion: restoration of PIG-A function produced no consistent effect on sensitivity to apoptosis [59]. If PIG-A gene product dysfunction rendered HSC resistant to apoptosis, expansion of the PNH clone would be due to an obvious intrinsic advantage. However, such a result would be inconsistent with the rarity of PNH, especially after mutagenic stress such as chemotherapy. Also lacking is a mechanism by which PIG-A mutations would somehow influence the apoptotic process, as none of the myriad members of the ever-expanding list of apoptosis-triggering cell surface molecules expressed by hematopoietic cells is known to be GPI-anchored. (There is, however, one recently reported example of an apparent GPI-anchored death transducer in nonhematopoietic cell lineages [60].) A recently described ‘‘decoy’’ death receptor with significant homology to the other members of the death receptor family, termed DcR1, has been found to be GPI-anchored [61], but loss of expression of DcR5 by a PIG-A mutant cell should increase susceptibility to apoptosis. Alternatively, an inactive PIG-A gene product might directly or indirectly affect biochemical pathways supplementary to the biosynthesis of the GPI anchor. The PIG-A gene product might participate in glycosylation of intracellular proteins involved in signal transduction analogous to the recently described posttranslational modification of the cytoplasmic estrogen receptor by covalent attachment of an


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N-acetyl-glucosamine [62]. For another example, ceramide is known to cotraffic with GPI-AP in polarized epithelial cells to the basolateral membrane [63]. If impaired synthesis of the GPI moiety were to result in altered ceramide trafficking, then membrane ceramide content would be diminished in PIG-A mutant cells, influencing signal transduction [64–69]. Apoptosis resistance of primary cells from PNH patients has been investigated [58, 59, 70]. While neutrophils and more primitive (CD34⫹) hematopoietic cells from PNH patients undergo less apoptosis than similar cells from healthy individuals, it does not appear that such resistance can be attributed to PIG-A dysfunction. The degree of resistance to apoptosis of neutrophils from different PNH patients did not correlate with the proportion of GPI-deficient neutrophils [59, 70], thus, two PNH patients with, for example, 10% vs. 90% GPI-AP(⫺) granulocytes may exhibit equivalent degrees of resistance to apoptosis compared to controls. Second, the apoptosis resistance observed for neutrophils and CD34⫹ cells from PNH patients is also seen in AA and MDS patients without PNH [70]. In vitro apoptosis resistance of hematopoietic cells from bone marrow failure patients may reflect an underlying pathologic process common to PNH, AA, and MDS. Resistance to apoptosis could be secondary to an accelerated cell cycle or to appropriate compensatory elevations in plasma concentrations of hematopoietic growth factors. Regardless of the precise mechanism, it is likely that only the most resistant cells of the normal Gaussian distribution of apoptosis resistance manage to survive under the conditions that produce bone marrow failure.

Studies in Genetically Engineered Murine Models Cloning of the PIG-A gene made possible the development of knock-out murine models, in which the direct effects of PIG-A gene product dysfunction on hematopoiesis in vivo could be studied in isolation. (See Chapter 6 by Kinoshita et al.) The uniform conclusion from studies of knock-out chimeric murine hematopoiesis is that a nonfunctioning PIG-A gene confers no intrinsic advantage to affected hematopoietic clones under physiological conditions [71–73], and instead, chimeric mice tend to show a trend of decreasing PIG-A mutant hematopoietic contribution over months of followup [73]. In vivo murine studies were initially performed with C57BL/6 blastocysts injected with 129/Sv PIG-A knock-out (KO) embryonic stem cells (ESC); thus, the mutant and wild-type contributions differed by more than the just the targeted disruption in PIG-A (germline incorporation of unconditional PIG-A mutations proved fatal in utero, thus preventing generation of an inbred PIG-A knock-out mouse strain). One of the initial eight PIG-A KO chimeric mice did develop a delayed dramatic expansion of GPIAP(⫺) erythrocytes, at age 10 months, and eventually died of unknown causes [71]. Whether this isolated instance of clonal expansion was related to the disrupted PIG-A gene or due to either an acquired mutation in the knock-out ESC line or the resultant mouse is unknown. Subsequent studies have therefore utilized F1 animals derived from inbred mice with modified PIG-A genes flanked by lox recombination sites crossed with mice transgenic for the Cre recombinase [73]. Again, no tendency for mutant HSC to expand in such systems was observed [74, 75]: The PIG-A-mutant contribution to the circulating red and white cell

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populations diminished gradually over time [74]. Caution should be exercised in extrapolating these results from mice to humans; the homologue(s) of a given human GPI-AP may not, in fact, be GPI-anchored in the mouse or such homologues may be GPI-anchored in both species but subserve fundamentally different physiologic (or pathologic) functions in humans vs. mice. For example, expression of CD24 (a GPI-anchored protein in both species) is largely restricted in humans to lymphocytes, while in mice it is present on diverse lineages, ranging from embryonic stem cells to terminally differentiated erythrocytes. It remains to be seen in knock-out mice whether, under conditions of experimental hematopoietic stress, PIG-A-mutant hematopoiesis is favored.

An Immune Model of PNH Pathogenesis To be consistent with the observations presented above, a model of PNH pathogenesis must posit both PIG-A mutations in the HSC pool and extrinsic selection of HSC which is dependent, at least in part, on some aspect of normal PIG-A function, most probably GPI-AP biosynthesis. Genetic stability in PNH patients may be normal [76] or increased as a presumed direct result of the underlying attack on the bone marrow [77]. We have estimated a 10⫺6 mutation rate in the hypoxanthine-guanine phosphoribosyl transferase (hgpt) gene in normal clonogenic progenitors (unpublished observations.) Using this conservative figure and an estimate of the number of human HSC (extrapolated from mice) of ⱖ300,000, an average human would have greater than a one in four chance of harboring at least one PIG-A mutant HSC. Empirical evidence of PIG-A mutations in mature myeloid cells has indeed been reported in healthy humans [78]. Araten and Luzzatto sorted CD55(⫺), CD59(⫺) granulocytes (defined by granularity and expression of CD11b) from healthy individuals and in over half of the subjects obtained mutant PIG-A sequences that were considered likely to result in an inactive gene product. In one person, the mutation introduced a new restriction endonuclease site permiting independent confirmation of the PIG-Amutant clone 5 months later by repeat sorting followed by gene amplification and restriction endonuclease digestion. The range of frequency of CD55(⫺), CD59(⫺) granulocytes in five normal subjects was 10 to 60 per million. While this study could not directly address whether the PIG-A mutation was present in the HSC pool, it is likely that at least some of the mutant cells arose from mutant primitive cells. Thus most healthy individuals appear to harbor minor PIG-A mutant hematopoietic clones. The precise mechanism by which profound selection of PIG-A mutant precursors takes place in PNH is unknown. Iatrogenic immunoselection of PIG-A mutant lymphocytes in humans has been documented by clinical investigators under artificial circumstances involving medical therapy [79–81]. Each group studied the unintended consequences of administering CAMPATH-1H (a humanized monoclonal antibody to CDw52, which is coincidentally GPI-anchored) to patients with either non-Hodgkin lymphoma, rheumatoid arthritis, or chronic lymphocytic lymphoma. Patients within each of these disease categories were observed to develop a major population of GPI-AP(⫺) lymphocytes after several courses of CAMPATH-1H and over a relatively short time period of 1 to 2


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months. The basis for the GPI-AP(⫺) phenotype in the emergent lymphocyte subpopulation was shown in several cases to be due to PIG-A mutations. Thus immune-mediated selection, here humoral, was adequate to drive expansion of variant cells that are negative for the targeted epitope; in the case of GPIanchored targets, mutations in PIG-A would appear to be the genetic events most likely to accomplish this. No case of AA or PNH, however, has ever been attributed to autoantibodies against GPI-AP on HSC. It has been proposed that the mere stress of repopulating a depleted (aplastic) bone marrow by a handful of surviving HSC may be sufficient pressure to select for the PIG-A mutant genotype. In order to address whether nonspecific myelosuppression is sufficient to select for significant expansion of PIG-A mutant progenitors, we examined GPI-AP expression in peripheral blood samples from several cohorts of patients whose hematopoietic progenitor pool had undergone contraction as a result of medical therapy [82]. PNH did not develop in either patients who had received multiple cycles of cytotoxic chemotherapy (N ⫽ 18), nor in patients who had undergone allogeneic BMT (N ⫽ 28). These figures are very different from the frequency at which PNH is observed in bone marrow failure patients: 25 of 115 AA patients and 9 of 39 MDS patients seen at the NIH (22%, P ⬍ 0.025). Thus, the mechanism of myelosuppression would appear to be critical. The clinical association of the rare diseases of PNH and AA (or MDS) is clearly an important clue to the underlying pathologic process. Collective clinical and laboratory data strongly implicate an (auto-) immune pathogenesis in AA. An activated CD8⫹ T cell appears to be the proximal cell responsible for the destruction of primitive hematopoietic cells [83]. Laboratory studies show that multiple parameters of T cell activation are elevated in AA patients (IFN-웂 mRNA expression in BM[84]; HLA-DR and IL-2 receptor expression by CD8⫹ cells [85]), and bone marrow function in patients improves after immunosuppression [86, 87] (see section on therapy). Moreover, syngeneic bone marrow transplants (using an identical twin) for AA will generally fail unless preceded by immunosuppressive conditioning [31, 32]. To account for the association of PNH with AA, several mechanisms can be hypothesized in which GPI-AP-deficient HSC might escape cytotoxic attack: A peptide derived from a GPI-anchored protein expressed by stem cells may serve as an autoantigen in some or all cases of AA, or perhaps only in those that progress to PNH (Fig. 5-2). However, because both GPI(⫹) and GPI(⫺) cells synthesize the polypeptide precursors [p(GPI-AP)TM] of mature GPI-AP[88], it would be necessary to postulate that in PIG-A mutant cells the proper (auto-) antigenic peptide fragment is inefficiently generated or mal-processed (see Fig. 5-2). Receptors on HSC for secreted CD8⫹ T cell products might be GPI-anchored (Fig. 5-3). However, no such examples are known and GPI-APs play no role in the action of IFN-웂, TNF-움, perforin, granzymes, or Fas ligand. (A GPI-anchored TGF-웁-binding protein of unknown signal-transducing capacity has been described in endometrium and in keratinocytes [89–91].) GPI-AP can transduce signals to the interior of the cell, at least in response to cross-linking by antibodies [92]. Less common are examples of natural

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Figure 5-2 PIG-A mutation and antigenic peptide presentation. A mature GPI-AP is generated in a transamidation reaction when the first 웆 amino acids (after the signal peptide) of the transmembrane precursor polypeptide of a GPI-AP [p(GPI-AP)TM] are transferred to the presynthesized GPI anchor. If no fully assembled GPI anchor is available to accept aa웆 in the transamidation reaction, then p(GPIAP)TM will be shunted to an ill-defined proteolytic scavenge pathway [88]. It is possible that cleavage through this default pathway might prevent generation of a full-length (putative) autoantigenic peptide derived from a properly pocessed GPI-AP species. Thus, if a peptide from a GPI-AP was an HSC autoantigen in AA, PIG-A mutant HSC would enjoy a survival advantage. (see also Fig. 5-1). , proteasome degradation sites; , default proteolytic cleavage sites for scavenging malprocessed p(GPI-AP); 䊉, normal site of transfer of p(GPI-AP) to presynthesized GPI anchor in the transamidation reaction; Ino; inositol.

ligands which employ GPI-anchored receptors, on any cell type, for signal transduction. (See Chapter 8 by Schofield.) The three best characterized of such ligands are ciliated neurotrophic factor (CNTF), neurturin, and glial cell line-derived neurotrophic factor. Intracellular signals of all three of these factors are transduced by heterodimers consisting of a factor-specific, GPIanchored receptor coupled noncovalently with a transmembrane partner [93–100]. While the target tissue of all three is neural, it is interesting to note that HSC express gp130 (the partner for the GPI-anchored CNTF receptor) as a heterodimer with either the IL-6 or LIF receptor (neither of which is GPI-anchored). Thus, if some undiscovered GPI-anchored partner


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Figure 5-3 Model in which a GPI-AP served as a receptor for an inhibitory ligand. (Top) A GPI-AP may serve as a receptor (or coreceptor) for a cytokine such as TGF-웁 or another, uncharacterized factor. Signal transduction by the GPI-anchored receptor may require cis-interaction with a transmembrane coreceptor, depicted as a rectangle (analogous to the interaction in neural cells between gp130, which is a transmembrane protein, and the receptor for ciliated neurotrophic factor (CNTFRc), which is GPI-anchored). (Bottom) The external inhibitory signal need not be soluble but may be cell associated. The cell delivering the negative signal could be a stromal cell, lymphocyte (see also Fig. 5-5), or some other cell.

for gp130 is expressed by HSC, then inability to respond to its ligand could potentially alter the behavior of such a PIG-A-mutant stem cell and confer a growth advantage. A cytopathic stem cell-tropic virus (Fig. 5-4, top) may either usurp a GPIAP for entry into HSC (nonstem cell-tropic examples include ECHO and coxsackie viruses [101–103]) or a noncytopathic virus may encode for a viral GPI-AP[104, 105] (vGPI-AP) that proves to be the immunodominant antigen (Fig. 5-4, bottom). In the latter case, PIG-A mutant HSC would fail to elicit what ultimately would be a deleterious immune response, and their expansion would thus be favored. A cytopathic viral model, however, would not account for the long-term clinical improvements frequently observed after treatment of uncomplicated AA with immunosuppression or BMT.

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Figure 5-4 Two viral models for selection of PIG-A mutant HSC. (Top) The viral receptor is GPIanchored. (Bottom) The viral receptor is not GPI-anchored, but the virus encodes for a GPI-anchoredmolecule, which happens to be the dominant antigen against which the immune response is directed. This latter model would allow for survival (and clonal expansion) of the infected HSC only in the event that the virus was noncytopathic.

Expression of one or more GPI-AP by the HSC may be necessary for optimal attack on wild-type HSC by the T cells that mediate AA (Fig. 5-5). Plausible mechanisms that could account for such targeting would include Some direct or indirect consequence of PIG-A mutations attenuates the transduction of apoptosis signals to HSC by immune effectors. (See above discussion and also the bottom of Fig. 5-5.) A GPI-anchored member of the ‘‘costimulatory’’ family [106–113] of molecules might exist (Fig. 5-5, top) and be expressed on HSC in either health or disease. Such molecules, however, have generally been described to be important primarily in the afferent phase of an immune response, and not in the effector phase. Thus, after establishment of autoimmunity in AA, which presumably develops at a time when a


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Figure 5-5 T lymphocyte-mediated selection of PIG-A mutant HSC. T cell receptor (TCR) engagement by an antigenic complex of (auto-) peptide plus HLA is presumably necessary (but may not be sufficient) for T cell activation. (Top) Expression of an uncharacterized GPI-anchored protein on the target HSC may be necessary for delivery of the immunotoxic signal. A transmembrane cis-associating partner for the GPI-AP (shaded ellipse) may be required for transduction of the inhibitory signal. (Bottom) Complete activation of the pathogenic T cells in AA may depend on costimulation by an accessory molecule which, on HSC, may be GPI-anchored.

patient is ⬎ 99% GPI(⫹), both wild-type [GPI-AP(⫹)] and PIG-A mutant [GPI-AP(⫺)] HSC targets should be equally susceptible to attack by activated T cells, unless periodic ‘‘repriming’’ with signal 1 (antigen or autoantigen) plus signal 2 (costimulator) is necessary for the longterm maintenance of autoimmunity. GPI-anchored ligands for T cell surface adhesion molecules may play an important role in T cell attack on HSC (Fig. 5-5, bottom). Three GPI-AP, CD58 (LFA-3), CD48, and CD59, serve as ligands of varying avidity for CD2[114–116], a signal-transducing molecule present on the surface of all T cells (and NK cells). The functional importance of interactions between CD2 and any of these three ligands, however, has yet to be demonstrated. For example, using EBV-transformed LCL (which

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express CD48, CD58, and CD59) as immunologic targets, Hollander and Springer demonstrated that a GPI-AP(⫺) daughter line was equally susceptible to cell-mediated killing in three different assays of killing [117]: by classical cytolytic (CD8⫹) T lymphocytes (CTL), NK cells, and in antibody-dependent cellular cytotoxicity (ADCC). On the other hand, some in vitro data suggest that circulating lymphocytes from PNH patients may exert GPI-dependent selection (Fig. 5-6). These experiments originated from the observation that PIG-Amutant LCL are very difficult to obtain from EBV transformations of PBL from PNH patients, even in patients with readily demonstrable GPI-AP(⫺) B cell subpopulations in their blood. The EBV receptor on B cells, CD21, is not GPI-anchored. These findings suggested that PIGA-mutant LCL were actually at a disadvantage as measured by cell proliferation or viral transformation. Because almost everyone has been naturally exposed to EBV, and thus normal PBL contain a population of T cells capable of destroying EBV-transformed B cells, it is necessary, when establishing LCL lines, to include CsA in the primary culture of PBL with EBV supernatant. However, when CsA was deliberately omitted from EBV transformations of PNH PBL, allowing expression of endogenous anti-EBV immunity, a discreet population of PIG-A mutant LCL emerged (Fig. 5-6). This phenomenon has been seen in about half of the PNH patients whom we have examined. These results are consistent with a relative survival advantage for the GPI-AP(⫺) pheno-

Figure 5-6 Autologous T cells from PNH patients preferentially suppress EBV transformation/expansion of GPI-AP(⫹) B cell clones. The left-hand figure shows the result when an unfractionated peripheral blood mononuclear cell (PBMC) culture from a PNH patient is transformed with EBV, under standard conditions. CsA is included in such routine transformations in order to inhibit the EBV-immune T cells present in the PBL of 95% of the population, which would otherwise destroy nascent EBV-infected B cell clones in vitro. CD20 (abscissa) is a marker for B cells. CD59 (ordinate) is a GPI-anchored antigen present on cells of all hematopoietic lineages, including B cells. Essentially all of the CD20(⫹) cells are GPI-AP(⫹) in the transformation in which CsA was included (left). The reason why GPI-AP(⫺) B cells (from PNH patients) are not efficiently transformed under these standard conditions is not known. The CD20(⫺)/dim cells are T cells. The right-hand figure shows the result when PBMC from the same PNH patient are EBV transformed in the absence of CsA. A GPI-AP(⫺) B cell population (bottom right) has been selected, presumably by uninhibited T cells.


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type under certain circumstances of immune attack. Precisely which effector population mediates this GPI-dependent selection and which receptor–ligand interactions are involved have yet to be determined. These results can be reconciled with those of Hollander and Springer [117] by taking into account that the target of immune attack in our system is not a fully transformed latently infected B cell blast (which is known to be resistant to certain modes of programmed cell death [118]), but rather a B cell at an earlier stage of EBV infection, which may not yet have acquired the apoptosis-resistant phenotype of established LCL, and that effectors in our system were derived from PNH patients, who may harbor fundamentally skewed, deranged, or hyperactive immune effector populations. A working model accommodating much of the data discussed above can be proposed in which the PIG-A mutation protects affected HSC from the cytotoxic autoimmune attack that characterizes aplastic anemia (Fig. 5-7). Protection could be conferred by loss of expression of one or more GPI-AP by mutant HSC, as this is the only known molecular consequence of PIG-A gene dysfunction. Future efforts need to focus on identifying the precise GPI-anchored molecules on HSC involved, as well as their cognate ligands secreted by or expressed on the surface of pathogenic lymphocytes (Fig. 5-6). Characterization of these molecules should then facilitate development of novel and, presumably, more narrowly targeted therapeutic approaches than currently available, as discussed below.

Clinical Course and Treatment of Marrow Failure in PNH Natural History Understanding the natural history of PNH is necessary in order to counsel patients regarding the relative risks and benefits of various therapies. Three large-scale analyses of the clinical course and prognostic factors in PNH have been published, comprising a total of 460 patients [23, 42, 119]. All used the Ham test for definitive diagnosis. Median survival ranged from 10 to 15 years after diagnosis [23, 42, 119–121]; approximately one-quarter of patients survived 25 years. Symptoms attributable to PNH may, in retrospect, precede diagnosis by as long as a decade. Approximately one-third of deaths in the European studies were related to thrombotic complications [23, 119], compared to fewer than 10% in the Japanese experience [42]. Conversely, hemorrhage (from thrombocytopenia due to aplastic anemia) was the largest single cause of death among Japanese [42] (39%), but only 15–25% of Europeans died of this complication [23, 119]. Of those patients presenting with thrombosis, 60% died within 4 years [119]. Transformation to leukemia occurs in fewer than 3% of patients [23, 42, 119, 121]. A curious prognostic feature noted in the French study [119] was that an antecedent history of aplastic anemia was associated with a 2.5-fold lower relative risk of death.

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Figure 5-7 A model of immune selection favoring expansion of PIG-A mutant hematopoiesis. Multiple T cell clones may pathologically target normal HSC. If these T cells require target expression of GPI-anchored costimulatory adhesion or death signal-transducing molecules for optimal attack and/or sustained (poly-)clonal expansion, outgrowth of even rare PIG-A mutant HSC will occur. The self-renewal capacity of stem cells will permit nonneoplastic mutant clones to support hematopoiesis for prolonged periods.

The replacement of the Ham test with much more sensitive flow cytometric techniques recently introduced for the detection of GPI-AP will doubtless result in more diagnoses of PNH, in milder cases, at earlier time points, and in a broader spectrum of patients (for example, MDS[82]). The net impact on time of survival postdiagnosis is difficult to predict; lead-time bias should lengthen the apparent median survival time, while it is possible that increased representation of the previously occult MDS/PNH cases may actually shorten it.


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Therapies Amelioration of Impaired Hematopoiesis The overlap of PNH and AA comes to medical attention in three chronologically distinct fashions. First, a patient presenting with pancytopenia may have a Ham test or flow cytometry performed in the course of their initial evaluation and the diagnosis of AA and PNH will be concurrent. Second, a patient with predominantly hemolytic PNH may be found, after months or years of observation, to develop absolute reticulocytopenia, thrombocytopenia, and/or neutropenia (although even during the hemolytic phase of PNH, evidence of impaired hematopoiesis may be demonstrable in vitro[43–47]). Finally, an AA (or MDS) patient may be diagnosed with PNH by Ham test or flow cytometry months or years after presentation with marrow failure. Unfortunately, it is common for neither test to have been performed at the time of intial diagnosis with AA, and AA patients do not exhibit a positive Ham test if heavily transfused with normal red cells. Of note, our current experience with 35 prospectively followed AA patients is that all patients ultimately destined to develop PNH (N ⫽ 5) had flow cytometric evidence of GPI-AP(⫺) granulocytes at the time of initial diagnosis with AA[82]. In all cases, however, the therapeutic approach to hypoplastic PNH is essentially the same as for AA[122], regardless of the sequence of diagnoses (see below). The treatments for the life-threatening cytopenic complication of PNH include immunosuppression with antithymocyte globulin (ATG) and cyclosporine A (CsA), or bone marrow transplantation (BMT, see below). The response rates [123–126] reported for ATG or CsA are generally ⱖ50%; our own unpublished experience with ATG plus CsA indicates improvement in 80–90% of cases of AA/PNH and MDS/PNH [82]. ATG administration may have added toxicity in PNH patients. Intravascular immune complex formation between equine antibodies and the human target leukocytes against which the horse was immunized can result in complement activation and lysis of GPI-AP(⫺) red cells. In patients with a high percentage of PNH erythrocytes, fever, rigors, hypotension, massive hemolysis, and/or renal failure may occur during ATG infusion, analogous to a major transfusion reaction [125]. Intravenous steroids may temper this reaction, but are usually administered regardless, as prophylaxis against serum sickness [127]. Slowing the rate of infusion of the ATG may also lessen the intensity of the reaction. Aggressive hydration and transfusional support should be given during the infusion. Despite their salutary effects, however, immunosuppressive therapies do not eradicate the GPI-AP-deficient hematopoietic clones [128], nor does the proportion of normal hematopoiesis inevitably improve [129, 130]. Accordingly, even responding patients remain at risk for future episodes of hemolysis or thrombosis. Bone Marrow Transplantation (BMT) BMT offers the only curative therapy for PNH. Unfortunately, fully engrafted patients may develop acute or chronic graft-versus-host disease (GVHD) and attendant infections, and thus some ‘‘cured’’ patients eventually die not from PNH but from complications of the procedure. The decision to proceed with transplant, therefore, must involve careful weighing of the potential benefits and risks of an irreversible intervention.

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It is clear from rare cases of PNH in which a syngeneic (identical twin) donor was available that an aggressive preparative regimen is necessary for consistent, durable replacement of the patient’s mutant HSC [29–32] (see Table 5-1). Preparative regimens typically consist of high-dose cyclophosphamide plus additional immunosuppressive and myeloablative components such as procarbazine, busulfan, antithymocyte globulin, or total body irradiation [31, 36, 131, 132]. Relapse of PNH after allogeneic marrow transplantation preceded by cyclophosphamide plus busulfan [36] or cyclophosphamide alone [37] has been reported but is rare (relapse in one patient was associated with a course of IFN-움 therapy for hepatitis). Only two groups have published results with more than a few HLA-identical sibling bone marrow transplants in PNH [29, 36, 131], and in only the Seattle report is it clear that all of the PNH cases transplanted were reported. Six out of six (95% confidence interval: 54–100%) of the HLA-identical sibling (nontwin) transplants performed at Seattle were alive at 2 to 19 years followup, although one of these patients had biochemical evidence of PNH post-BMT, despite a preparative regimen which included busulfan [36]. Four of these six patients developed chronic graft-versus-host disease. The report from Boston [131] describes four transplanted PNH patients, all of whom were alive and well, with full donor chimerism, at the time of followup (1–60 months). One PNH patient treated with two consecutive haploidentical-related transplants died of pulmonary hemorrhage at 39 days after the first transplant without evidence of engraftment [36]. The current consensus indications for BMT in PNH are the development of either aplastic or thrombotic complications. Unfortunately, first thrombosis is often permanently disabling or fatal [23]. On the other hand, Budd-Chiari syndrome has been shown to resolve in at least one PNH case after BMT [132]. Hemolysis as the sole clinical manifestation of PNH is not considered sufficient indication to warrant BMT. Because children with PNH appear to have a worse prognosis [123], fare better the rigors of transplantation, and have a greater potential survival benefit, early transplant has been suggested in the pediatric population [123]. Future Therapies Additional therapies for PNH need to be investigated. The ‘‘mini-transplant,’’ consisting of a less toxic preparative regimen of fludarabine, high-dose cyclophosphamide, and ATG, may be found to provide a more favorable risk–benefit ratio than traditional BMT in PNH, in particular if a graft-vs.-autoimmunity effect exists. Alternatively, various biological response modifiers such as IL-10, CTLA4-Ig, humanized anti-CD25, soluble TNF-receptor, and mycophenolate mofetil might prove effective in reversing the pathology underlying the expansion of PIG-A mutant stem cell clones.

Summary PNH is an acquired clonal disorder in which PIG-A mutant clones of hematopoietic stem cells undergo massive but nonneoplastic expansion. The only known


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consequence of PIG-A gene dysfunction is loss of GPI-AP expression by the mutant HSC and its progeny. In erythrocytes, this defect results in episodes of intravascular hemolysis when complement is activated in the context of inflammation; in platelets, the defect is believed to favor thrombus formation. On the other hand, PIG-A mutations do not appear to directly give rise to the bone marrow aplasia that frequently accompanies PNH. Instead, it would seem that the underlying pathophysiologic process of AA confers a relative survival advantage to PIG-A mutant stem cells. Clarifying the molecular basis for this ‘‘unnatural’’ selection process will likely impact our understanding not only of PNH, AA, and other autoimmune syndromes, but also of the unique biology of GPIanchored proteins as well as the adaptive physiology of hematopoietic stem cells.

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6 Animal Models of PNH Taroh Kinoshita,* Monica Bessler,† and Junji Takeda** *Department of Immunoregulation, Research Institute for Microbial Diseases, Osaka University, Osaka 565-0871, Japan; †Division of Hematology, Department of Internal Medicine, Washington University School of Medicine, St. Louis, Missouri 63110-1093; and **Department of Environmental Medicine, Osaka University Medical School, Osaka 565-0871, Japan

Introduction As described in previous chapters, paroxysmal nocturnal hemoglobinuria (PNH) is a unique example of somatically acquired genetic disease. The responsible gene and its somatic mutations have been well characterized. A full understanding of mechanisms and events that lead to the clinical manifestation of PNH could establish a paradigm for the pathogenesis of acquired genetic diseases. It is easily conceivable that two events are required for the clinical manifestations of an acquired genetic diseases: (i) generation of mutant cell(s) through somatic mutation and (ii) clonal expansion of these cells. The latter is required because somatic mutation would occur in one or a few cells and for a clinical disease to develop, a significant number of abnormal cells must be generated. This is also true in particular for PNH. Analyses of PIGA mutations in affected blood cells from patients demonstrated that they harbor one to four mutant clones, one of them being a predominant clone in most cases [1–3]. The major clone occupies 10 to nearly 100% of myelopoiesis and also a significant percentage of lymphopoiesis (although the latter percentage is usually smaller than the former). The mechanism responsible for clonal expansion of PIGA mutant is not clear at the moment. One important point is whether mutation of PIGA alone confers hematopoietic stem cells with an ability to expand or whether an additional factor is involved. This issue can be addressed using animal models. Employing gene targeting technology, we generated Piga-disrupted embryonic stem (ES) cells and derived from them Piga-disrupted hematopoietic stem cells and PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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their progenies in vitro and in vivo. We describe these experiments here and discuss their impact on the understanding of the pathogenesis of PNH. In the following discussion, PIGA will refer to the human gene, Piga to the mouse gene, and PIGA to the gene products of both species [4].

Mouse Piga Gene Genetic Characteristics A cDNA of mouse Piga was cloned by means of colony hybridization with a human PIGA cDNA probe and 5⬘ and 3⬘ RACE methods [5]. A full-length mouse Piga cDNA contained 3557 bp, which is slightly shorter than that of human PIGA (3589 bp) (see below) [6]. The coding region of mouse Piga spans nucleotides 86 to 1540 and encodes a total of 485 amino acids; similarly, the coding region of human PIGA spans nucleotides 86 to 1537, encoding 484 amino acids. Thus, the mouse PIGA protein is only one amino acid longer than the human PIGA protein; they have 88% amino acid identity (Fig. 6-1). Transfection of the mouse Piga cDNA restored the surface expression of Thy-1 on BW5147 Thy-1⫺a mutant mouse lymphoma cells [5] which are defective in surface expression of this glycosylphosphatidylinositol (GPI)-anchored protein Thy-1 due to a defect in the first step of GPI-anchor biosynthesis [7, 8]. Moreover, Piga mRNA was completely missing in BW5147 Thy-1⫺a cells [5]. Therefore, Piga is the defective gene in this mutant, and the mouse PIGA is functionally homologous to human PIGA. Using a mouse Piga cDNA probe, genomic clones of Piga were isolated and analyzed [5]. The Piga gene spans 16–17 kb and has six exons (Fig. 6-2). Exon lengths range from 23 to 2281 bp, and their splice-site sequences are consistent with the AG/GC rule (Table 6-1). Exon 1 encodes a part of the 5⬘ untranslated region. Exon 2 encodes the rest of the 5⬘ untranslated region and the aminoterminal portion of PIGA protein. Exon 6 encodes the carboxyl-terminal portion of the protein and the 3⬘ untranslated region (Fig. 6-2). The structure of mouse Piga gene is very similar to that of the human PIGA gene [9, 10]. Mouse exons 1, 3, 4, and 5 are exactly the same sizes as their human counterparts. Mouse exon 2 is three nucleotides longer than human exon 2, making the mouse protein one amino acid longer than the human protein. The mouse Piga gene is about 1 kb shorter than human PIGA gene because of smaller introns. Finally, mouse Piga was mapped to the X chromosome at F3/4 [5]. Mouse XF3/4 is synthenic to human Xp22.1 where PIGA is localized [10, 11]. These results indicate that the mouse Piga gene has function, structure, and chromosome location similar to those of the human PIGA gene. Disruption of mouse Piga is, therefore, a good model system to study the pathogenesis of PNH in mice.

Figure 6-1 Comparison of deduced amino acid sequences of mouse and human PIGA and Saccharomyces cerevisiae SPT-14. The alignment was maximized by introducing the insertions, marked by dashes. Amino acids identical in at least two members are shown in the top line. Dashes in the top line indicate positions where only one member has residues. MPig-a and hPIG-A indicate mouse and human PIGA, respectively. (Reproduced from [5].)

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Figure 6-2 Structures of the human PIGA and mouse Piga genes. Boxes indicate exons; shaded areas are coding regions. Lines connecting exons indicate introns. The splicing patterns of human PIGA are indicated by thin lines. (Reproduced from Adv. Immunol. 60, 71 (1995).)

Piga Gene Inactivation in ES Cells ES cells used for homologous recombination of the Piga locus were male (female ES cells are genetically less stable and therefore less frequently used for in vitro gene targeting). A single recombination event involving the only Piga gene on the male X chromosome is therefore sufficient to abrogate the expression of glycosylphosphatidylinositol (GPI)-anchored proteins on the cell surface of ES cells. Kawagoe and colleagues [12] and Dunn and colleagues [13] disrupted the function of the Piga gene by replacing the 5⬘ end of Piga exon 2, encompassing the first ATG of the open reading frame, with the neomycin phosphotransferase gene under the control of the phosphoglycerate kinase promoter (PGK neo) (Fig. 6-3). In similar experiments, Rosti et al. replaced the 3⬘ end of Piga exon 2 with a selectable maker gene [14], based on the results of mutation analysis in patients with PNH, which had shown that the 3⬘ end of exon 2 is essential for the PIGA protein to form a functional glycosyltransferase complex [15]. In all these studies, the function of the Piga gene was completely abolished as shown by the lack of GPI-anchored proteins like heat-stable antigen (HSA or CD24) on TABLE 6-1 Sequences of Intron–Exon Boundaries in the Murine Pig-a Gene No.

Exon (bp)

Intron (kbp)

Donor

Acceptor

1 2 3 4 5 6

23 780 133 133 207 2281 ⫹ poly(A)

3 4.5 0.45 0.138 2.5

CTGGAGGgtaagta a AGAAAAGgtaatga ATGACAGgtattca TTTGCAGgtaaaac TGAAAAGgtaaggc

tctctttcagGTGACAG acaatttcagGGACTGA caaaaattagGGTGCAG ttttcttaagGTCGTCA tctcttgcagGTGTATG

a

Exon sequences are shown in capital letters and intro sequences in lower case letters.


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Figure 6-3 Homologous recombination of the Piga gene, (a) Genomic structure of the Piga gene. Black and white boxes represent noncoding and coding regions, respectively. Exon numbers are indicated below the boxes. Probes H and K were used for Southern blots to confirm the targeted disruption of Piga gene. Primers E and G used to detect the intact Piga gene are also indicated. (b) Piga targeting vector. The neo gene was inserted into exon 2 that contains a translational initiation site. The herpes simplex virus thymidine kinase gene was included at the 5⬘ end to select against random integration. (c) Structure of disrupted Piga gene after homologous recombination. Primers A and B were used to determine homologous recombination in ES cells. Primer F used with primer G to assess chimerism is indicated. Restriction enzyme sites: RI, EcoRI; B, BamHI; Sp, SpeI; Sa, SacI. (Reproduced from [12], by copyright permission of W. B. Saunders.)

the resulting ES cells. Piga-disrupted ES cells were used for in vitro differentiation experiments and in making chimeric mice.

In Vitro Differentiation of PIGA(ⴚ) ES Cells Background Most of what is known about hematopoiesis in PNH comes from clinical observations [16] or from in vitro culture studies of hematopoietic progenitor cells obtained from the bone marrow or peripheral blood of patients with PNH [17– 19]. One of the limitations of in vitro studies of hematopoiesis from patients with PNH is that progenitor number is significantly reduced [17, 19], making it difficult to investigate the functional effect of a mutated PIGA gene on the proliferation and differentiation of the hematopoietic progenitors and their progeny. The recently developed technology of targeted gene inactivation in murine ES cells is an ideal alternative approach to studying the consequences of a nonfunctional Piga gene on hematopoiesis and on other developmental programs. For a better understanding of the following sections and to highlight the implica-

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tion of hematopoietic differentiation of ES cells after genetic ablation of all GPIanchored proteins, we will first briefly review current knowledge of murine hematopoiesis in the developing embryo.

Genesis of Blood Cells in Embryo The developing hematopoietic system in mice is characterized by successive phases of hematopoietic stem cell differentiation (for review see Robb, 1997 [20] and Dzierzak et al., 1997 [21] and references therein). Figure 6-4 schematically summarizes the sequential waves of blood cell formation at different anatomical sites during mouse ontogeny. The first recognizable blood cells are primitive embryonic hematopoietic cells, which produce mainly nucleated erythrocytes expressing embryonic hemoglobin; primitive red cells appear in the blood cell islands of the extraembryonic yolk sac at about day 7.5 of gestation. When cultured in vitro, the yolk sac has been shown to contain not only erythroid progenitors but also precursors that have the potential to differentiate into cells of the granulocyte-macrophage lineage [22]. At about day 8.5, definitive hematopoiesis commences in the aorta-gonad-mesonephros (AGM) or paraaortic splanchnopleura region [23, 24]. Fetal hematopoiesis involves the differentiation of stem cells that have colonized the developing liver. Yolk sac progenitors are thought to contribute to the first wave of hematopoietic activity in the fetal liver at around day 9 followed by progenitors of definitive or adult hematopoiesis from the AGM region at day 10 (for review see [25] and references therein). In addition to the production of fetal, and later adult erythropoietic cells, hepatic hematopoiesis yields progenitors of both the myeloid and lymphoid lineages,

Figure 6-4 Genesis of murine blood cell production. Schematic representation of the successive waves of hematopoietic differentiation at different anatomical sites (see also text). The time scale in days of gestation (E ⫽ embryonic day) in which hematopoiesis is observed at specific anatomical sites is shown below. ES cells correspond to inner mass cells of a preimplantation embryo at E3.5-4.


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including for granulocytes, macrophages, B lymphocytes, and T cells. Around day 16, hematopoiesis shifts from the fetal liver to the spleen and to the developing bone marrow. The bone marrow is the main site of blood cell production in the adult mouse. The origin of embryonic and fetal hematopoietic stem cells (HSC) capable of long-term repopulation of irradiated mice is a subject of intense interest and some debate. HSC have been identified in the yolk sac at day 9, in the AGM region around day 8.5, and in the fetal liver [26, 27].

Embryonic Stem Cell Differentiation in Vitro ES cells are derived from the inner cell mass of the preimplantation embryo (for review see Robertson 1987 [28] and references therein). ES cells cultured in vitro on embryonic fibroblasts or in the presence of leukemia inhibitory factor (LIF) remain totipotent. When implanted into a host blastocyst, ES cells are competent to contribute to all tissues including germ line cells [29]. When removed from contact with feeder cells, or from the presence of LIF, and cultured in liquid or methylcellulose-containing media in bacterial grade dishes, ES cells, unable to adhere to the surface, generate colonies of differentiating cells known as embryoid bodies (EBs). Cell differentiation in EBs occurs in a temporally and spatially defined pattern that recapitulates the initial stages of embryonic development (for review see Keller 1995 [30] and references therein). Likewise, hematopoiesis in differentiating ES cell cultures parallels embryonic events [31] (see also Fig 6-4): the in vitro recapitulation of hematopoiesis during embryonic development allows access to populations of early precursors that are difficult, if not impossible to sample in vivo. For instance, studies with ES cells have provided insights into the controversial origin of HSC. ES cells differentiating in vitro from blastlike cells, when cultured as clones in the presence of vascular endothelial growth factor (VEGF) and c-Kit ligand, produce single colonies that contain primitive erythrocytes and multiple definitive blood cell lineages. These findings suggest that during embryogenesis there is a population of common progenitors that can give rise to stem cells, which are subsequently committed to differentiate along either primitive or definitive lineages [32]. More recent studies have shown that these blast colony-forming cells not only give rise to primitive and definitive hematopoietic progenitors but also to endothelial precursors, implying that this cell population represents the previously hypothesized hemangioblast, the common precursor of the hematopoietic and endothelial lineages [33]. In vitro differentiation of mutant murine ES cells obtained by gene targeting through homologous recombination yields valuable information that complements and extends gene knockout studies in animals. The in vitro system can be used to establish structure/function relationships for proteins involved in cell differentiation in a physiological context, and thus is superior to studies of heterologous cell lines. An excellent classic example is the use of the murine ES system in studies of GATA-1 mutant hematopoiesis [34–37]. Differentiation of ES cells can be achieved by different methods, which as applied to in vitro differentiation of PIGA(⫺) ES cells are summarized in Fig. 6-5. Dunn et al. [13] used standard liquid differentiation culture in bacterial grade petri dishes while Rosti and

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Figure 6-5 Culture systems used for differentiating PIGA(⫺) ES cells in vitro. (a) ES cells are induced to form EBs in methylcellulose and (b) in liquid culture. Hematopoietic differentiation is identified: (1) By microscopic examination of EBs for the presence of hematopoietic cells (hemoglobinized erythroid cells); (2) examination of May-Gru¨nwaldGiemsa stained cytospin preparations obtained from disaggregates EBs; (3) hematopoietic colony formation of differentiated ES cells after dissagregation of EBs and secondary plating into methylcellulose; (4) PCR or RT-PCR to confirm the presence of mutated Piga gene in hematopoietic colonies and to assess the expression of lineagespecific mRNA.

colleagues plated recombined ES in methylcellulose-containing media in petri dishes [14]. Hematopoietic differentiation was studied by visual inspection of EBs for the presence of hemoglobinized erythroid cells and/or by secondary plating of the differentiated EBs cells into methylcellulose-containing media supplemented with growth factors that support hematopoietic differentiation.

PIGA(⫺) Embryonic Stem Cell Differentiation PIGA(⫺) Cells Form Few and Structurally Abnormal EBs Results from several studies show that PIGA(⫺) ES cells form fewer and smaller EBs compared to PIGA(⫹) ES cells [13, 14]. The small PIGA(⫺) EBs also failed to show formation of a central cystic cavity which normally develops over the course of several days in suspension cultures by PIGA(⫹) EBs [38, 39]. These findings suggest that differentiation of PIGA(⫺) ES cells under these conditions is altered and is unable to undergo the usual structural arrangements found in EBs formed by PIGA(⫹) ES cells.


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PIGA(⫺) EBs Grown in Suspension Cultures Do Not Show Hematopoietic Differentiation Using the liquid culture technique to differentiate PIGA(⫺) ES cells in vitro, Dunn and colleagues found no evidence of hematopoietic differentiation in PIGA(⫺) EBs. Enzymatic disaggregation of the EBs followed by secondary plating into methylcellulose-containing media enriched with hematopoietic growth factors failed to generate morphologically recognizable hematopoietic colonies. However, coculture of PIGA(⫺) ES cells with PIGA(⫹) ES cells for 3–5 days partially restored the capability of the PIGA(⫺) ES cells to undergo hematopoietic differentiation. PIGA(⫺) ES cells expressed low levels of the GPI-anchored protein CD24 after cocultivation with PIGA(⫹) ES cells, suggesting that cell-to-cell transfer of GPI-anchored proteins had occurred during the period of coculture. Cell-to-cell transfer of GPI-anchored proteins has been previously demonstrated [40–42]. The authors therefore concluded that PIGA(⫺) EBs formed in suspension culture are unable to undergo hematopoietic differentiation, but that hematopoietic differentiation could be restored after cell-to-cell transfer of GPI-anchored proteins.

PIGA(⫺) EBs Grown in Culture in Methylcellulose-Containing Media Show Hematopoietic Differentiation Rosti and colleagues plated wild-type and mutant ES cells directly into methylcellulose. As in suspension culture, fewer and smaller EBs were formed when compared to the EBs formed by the PIGA(⫹) parent ES cells [14]. The number of EBs formed by PIGA(⫺) ES cells was partially restored when, instead of single cells, aggregates of PIGA(⫺) ES cells were seeded into the methylcellulose-containing media. An increase in the formation of EBs in methylcellulose by plating of cell aggregates had been described previously [43], suggesting that cell-to-cell contact or para and/or autocrine factors enhance the formation of EBs. When cultured in methylcellulose according to these results, PIGA(⫺) cells in EBs were competent for hematopoietic differentiation. Although the proportion of EBs with hematopoietic differentiation was lower compared with EBs derived from wild-type ES cells, microscopic examination clearly demonstrated the presence of hemoglobinized erythroid cells (see Fig. 6-6). In vitro progenitor colony assay of disaggregated PIGA(⫺) EBs showed the formation of hematopoietic colonies, confirming that GPI-anchored proteins are not essential for hematopoietic differentiation. At first sight, the findings by Rosti and colleagues seem to be in contrast with the data reported by Dunn et al. An interesting possibility that may help to explain this discrepancy is that correct spatial organization is required for formation of blood cell island in EBs formed in suspension cultures, but is less important in the compact growth condition of EBs in methylcellulose. Indeed a similar discrepancy in the capability of hematopoietic differentiation of EBs formed in suspension cultures and EBs grown in methylcellulose has been reported previously in differentiating GATA4(⫺/⫺) ES cells. GATA 4(⫺/⫺) EBs do not form a visceral endoderm and, similar to PIGA(⫺) EBs, lack a cystic cavity when cultured in suspension [44]. That the correct spatial organization is required for hematopoietic differentiation might also be an alternative explanation as to why hematopoietic differentiation was restored when PIGA(⫺) cells were transiently cocultured with PIGA(⫹) ES cells.

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Figure 6-6 Hematopoietic differentiation of PIGA(⫺) EBs obtained in methylcellulose cultures. EBs derived from wild type (A) and from PIGA(⫺) (B) ES cells after 8 days in methylcellulose culture. The arrows show EBs with hematopoietic differentiation. Note smaller size of EBs in B (original magnification ⫻100). Hematopoietic differentiation in EBs derived from wild type (C) and from PIGA(⫺) (D) ES cells (original magnification ⫻600). Cytospin preparation from wild type (E) and PIGA(⫺) (F) with hematopoietic differentiation: erythroid and myeloid cells are present both in wild type and PIGA(⫺) u. Staining with May-Gru¨nwald-Giemsa or Wright, respectively (original magnification ⫻400). (Reproduced from [14] by copyright permission of The American Society for Clinical Investigation.)


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Conclusion ES cells that lack expression of GPI-anchored proteins due to a mutation in the Piga gene are able to undergo hematopoietic differentiation in vitro. In vitro differentiation experiments further show that hematopoiesis can occur even in an environment that lacks GPI-anchored molecules, indicating that Piga is not essential for hematopoietic differentiation. However, no growth advantage in favor of the mutated cells was found in either culture system, suggesting that for the clonal dominance of PIGA(⫺) blood cells seen in patients with PNH, additional factor(s) are needed. The ability to differentiate PIGA(⫺) cells in vitro enables us to study PIGA(⫺) hematopoietic precursors, which might be difficult to access otherwise.

Chimeric Mice Bearing PIGA(ⴚ) Hematopoietic Stem Cells Generation of Chimeric Mice with PIGA(⫺) ES Cells Kawagoe and colleagues [12], and Rosti and colleagues [14] generated chimeric mice by injecting Piga-disrupted male ES cells into blastocysts or by aggregating them with 8-cell stage embryos. As described above, the Piga-disrupted male ES cells had no surface expression of GPI-anchored proteins. These ES cells had a lower ability to contribute to hematopoiesis than PIGA-competent ES cells: only mice with low levels of chimerism were born [12, 14]. This phenotype was not caused by an artificial factor(s) such as the use of inappropriate ES cells, because Piga-disrupted ES cells rescued with a Piga cDNA gave rise to mice with a high level of chimerism [12]. Some chimeric fetuses had higher numbers of Piga negative erythrocytes but simultaneously showed abnormalities of visceral organs which were not compatible with life [14]. Therefore, the relatively low contribution of Piga-disrupted ES cells to hematopoiesis was probably not due to an intrinsic inability of PIGA(⫺) ES cells to differentiate into cells of hematopoietic lineages. These data are in contrast to observations of hematopoiesis in mice chimeric for cells with a disrupted GATA-1 gene, another X-linked gene [45]: although these mice lack ES-derived erythroid cells, indicative of such an essential role of GATA-1 in erythroid development, the contribution to other organs was relatively high. GATA-1 is involved in maturation of blood cells within the erythroid and megakaryocytic lineages, and is also expressed in Sertoli cells of the testis, but it is not essential for the development of nonhematopoietic tissues. Data from mice chimeric for PIGA(⫺) cells therefore suggest that GPIanchored proteins have a crucial role in mouse development but not in hematopoiesis. A lower contribution of PIGA-deficient ES cells in chimeric mice would be lethal to the embryo, explaining the survival of mice with only a low level of chimerism [12, 14]. Among these animals, only a few had PIGA(⫺) hematopoietic cells. PIGAdeficient ES cells contributed to all blood cell lineages, consistent with the in vitro experiments described above. However, lineage preference was observed; PIGA-deficient erythrocytes were always detected in hematopoietic chimeras,

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while PIGA-deficient lymphocytes and granulocytes were detected much less frequently [12, 14].

Piga Disruption Alone Is Not Sufficient for Clonal Dominance In order to determine whether Piga disruption is sufficient to cause clonal expansion of PIGA-deficient clone(s), the proportion of GPI-negative erythrocytes in peripheral blood of these hematopoietic chimeras was monitored [12, 14]. The percentage of PIGA(⫺) blood cells was stable or even decreased (Fig. 6-7) [12, 14], suggesting that disruption of the Piga gene was insufficient to cause a clonal expansion of PIGA(⫺) blood cells and that additional factor(s) are involved in the cause of dominance of PIGA(⫺) cells in patients with PNH. However, one of the chimeric mice studied by Kawagoe and colleagues, which contained the highest percentage of GPI-anchor negative erythrocytes in the periphery, showed a sudden increase of PIGA(⫺) erythrocytes 1 year after birth [12]. The increase of PIGA(⫺) cells was also observed in the myeloid and lymphoid lineages, indicating that a clonal expansion had occurred at the level of hematopoietic stem cells, as in PNH [12]. The increase of PNH hematopoiesis in this single animal may be interpreted in two ways: (i) a second rare event not related to Piga disruption occurred in this particular mouse or (ii) a second consequence of Piga disruption led to the later increase of GPI-negative cells. If the latter were true, one would anticipate that an increase of PIGA(⫺) cells would be observed after a latent period in most chimeric mice that have high percentage of GPI-anchor negative erythrocytes.

Figure 6-7 GPI-anchor deficient erythrocytes in Piga-disrupted chimeric mice. The percentages of GPI-negative erythrocytes were monitored from 3 weeks after birth. 38 and 88 represent ES clone numbers. ICR in parentheses shows that ES cells (clone 38 or 88) were aggregated with 8-cell-stage embryos from ICR mice. BL/6 in parentheses shows that ES cells (clone 88) were injected into blastocysts from C57BL/6 mice. Mice a, b, c, and f died at 17, 10, 5, and 5 months of age, respectively. (Reproduced from [12] by copyright permission of W. B. Saunders Co.)


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Clonal competition of hematopoietic stem cells in chimeric mice generated from PIGA(⫺) ES cells is different from that which occurs in patients with PNH. Figure 6-8 compares the genetic background of HSC in chimeric mice with the situation present in patients with PNH. Because PNH is an acquired disease, the genetic background of PIGA(⫹) and PIGA(⫺) HSCs is the same. In chimeric mice PIGA(⫺) HSCs developed from PIGA(⫺) ES cells were derived from the 129 mouse strain, whereas PIGA(⫹) HSCs were of host origin (C57BL/6 or ICR), a difference that might be relevant for the clonal expansion found in one of the chimeric mice. The problem of the genetic background and the embryonic lethality of highly chimeric mice can be minimized by the use of conditional gene targeting.

Conditional Piga Disruption by Means of Cre/loxP System The relationship between deficiency of GPI-anchored proteins and high embryonic lethality was further studied using a Cre/loxP conditional gene knock-out technology [46]. For this purpose, ES cells bearing insertions of loxP sites in the flanking regions of exon 6 of Piga (Piga flox) were first generated by means of conventional gene targeting (Fig. 6-9A) [47]. The resulting Piga flox ES cells normally expressed GPI-anchored proteins (Fig. 6-9B, left panel). After transfection of Cre-expression plasmids into the Piga flox ES cells, the surface expression of GPI-anchored proteins was completely lost (Fig. 6-9B, right panel), indicating that Piga flox gene was efficiently disrupted due to a deletion of exon 6 caused by expressed Cre recombinase. Mice bearing Piga flox allele (Piga flox mice) were then generated using Piga flox ES cells (Fig. 6-9C). To disrupt Piga in the intact animal, Piga flox mice were mated with hCMVCre transgenic mice [48], which express Cre recombinase before transplantation by the action of human cytomegalovirus promoter. In male embryos that received

Figure 6-8 Comparison of the genetic background between chimeric mice generated from PIGA(⫺) ES cells and in patients with PNH. Schematic representation of hematopoietic and nonhematopoietic pools in chimeric mice and in PNH patients. In PNH, GPI-anchor negative cells are present only in the hematopoietic cell pool. In chimeric mice, GPI-anchor negative cells are present not only in the hematopoietic cell pool but in other tissues. Another critical difference in chimeric mice is that genetic background of GPI-anchor deficient and normal cells is different.

Figure 6-9 Targeted insertion of loxP sites in the Piga gene. (A) Part of the wild-type Piga locus containing exons 3, 4, 5, and 6, the targeting construct, and Piga allele inserted with loxP sites, are shown. Closed and open boxes are coding and noncoding exons, respectively. Restriction sites of BamHI (B) and EcoRI (E) are indicated. (B) Cremediated disappearance of GPI-anchored proteins on the ES cells containing loxP sites in Piga gene. The plasmid pMC-Cre16 with hygromycin resistance gene (hygr) was transfected into clone 58 of ES cells bearing Piga flox. The surface expression of heat-stable antigens (HSAg) was examined before and after selection with 150 애g/ml of hygromycin B for 7 days. Five of 12 clones lost surface GPI-anchored proteins after selection. A representative clone showing absent GPI-anchored proteins is illustrated. (C) Southern hybridization of DNA from mice bearing Pigaflox and control mice. Genomic DNAs from the tail were obtained and digested with BamHI. The 3.5 and a 2.0 kb bands represent targeted and endogenous alleles, respectively. (Reproduced from [47], copyright 1997 National Academy of Sciences, U.S.A.)


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an hCMV-Cre transgene and an X chromosome bearing Piga flox gene, complete bodywide disruption of Piga occurred, and development halted at day 9 of gestation due to totally impaired organ formation [46]. This lethal phenotype provides formal proof that GPI-anchored proteins have vital roles in embryonic development and some are likely involved in morphogenesis. In female embryos that received an hCMV-Cre transgene and one X chromosome bearing the Piga flox gene, one of the two Piga genes was disrupted widely in all tissues. The result was a mosaic PIGA-deficiency phenotype due to random inactivation of one of the X chromosomes through the process of Lyonization: cells in which the X chromosome bearing the disrupted Piga was inactivated became PIGA positive and GPI positive, and cells in which the normal X chromosome was inactivated became PIGA negative, GPI negative. The mosaic, partial PIGA deficiency also caused developmental abnormalities, especially in the cephalic regions [46]. The mice survived until the neonatal stage [46]. The roles of GPI-anchored proteins in specific organs were also investigated using Piga flox mice. Piga flox mice were mated with Keratin 5-Cre mice (K5-Cre) which express Cre recomibinase exclusively in skin [47]. In male F1 mice that received the Piga flox gene and the K5-Cre transgene, complete disruption of Piga occurred in keratinocytes, resulting in a nearly complete lack of GPI-anchored proteins in skin. These mice had abnormal differentiation of the cutaneous horny layer and died within few days of birth [47]. GPI-anchored proteins thus appear to be essential in skin development. Similar experiments were done using Lck-Cre transgenic mice that express Cre in immature T lymphocytes. In male mice which received both Lck-Cre transgene and Piga flox, efficient disruption of Piga occurred in thymic immature T lymphocytes. Due to the slow turnover rates of surface GPI-anchored proteins, residual Thy-1 and Ly-6 expression was detected in the thymus, making it difficult to evaluate the function of GPI-anchored proteins in T lymphocyte development. Nevertheless, GPI-anchored protein surface expression was completely lost in mature T lymphocytes of spleen and lymph nodes [49]—surprisingly, GPI-anchor deficient T lymphocytes behaved equivalently to GPI-anchor sufficient T lymphocytes in vivo and in vitro [49]. Cre transgenic mice should also be useful to examine the functions of GPIanchored proteins in other tissues and organs in vivo.

Generation of Bone Marrow Chimeric Mice Bearing PIGA(⫺) Hematopoietic Progenitors To circumvent the problems found in chimeric mice generated with PIGA(⫺) ES cells, bone marrow chimeric mice were generated taking advantage of the Cre/loxP system. The production of bone marrow chimeric mice relied on the genetically manipulated mice bearing Piga flox and hCMV-Cre, as described above. In Piga flox and hCMV-Cre double positive female mice, mosaic Piga deficiency occurred due to random inactivation of the X chromosomes. Because they were viable until day 20 of gestation, HSCs were obtained from day 14 fetal livers. Transfer of those HSCs into lethally irradiated mice allowed reproducible production of bone marrow chimeric mice. In these bone marrow chimeric mice, both PIGA(⫹) and PIGA(⫺) hematopoietic stem cells were of the same donor

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Figure 6-10 Bone marrow chimeric mice generated by using Cre/loxP system. Generation of bone marrow chimeric mice. Because fetal liver cells from Piga flox and hCMV-Cre double positive female mice contained both GPI-anchor deficient and normal hematopoietic progenitors, they were transferred into lethally irradiated host mice, C57BL/6, in order to generate bone marrow chimeric mice. The mice bore GPI-anchor deficient cells only in the hematopoietic cell pool, where GPI-anchor sufficient cells (wild type) were also derived from the same genetic background.

origin (Fig. 6-10). Therefore, these bone marrow chimeric mice resemble patients with PNH in terms of genetic background. Monitoring of GPI-anchor negative erythrocytes in these bone marrow chimeric mice is underway.

Hematopoiesis by Cells from Patients with PNH in SCID Mice As discussed above, studies with model mice generated by means of targeting disruption of the Piga gene have suggested that Piga-disrupted, GPI-negative hematopoietic stem cells do not have an intrinsic growth-promoting phenotype. However, when Nakakuma and colleagues transplanted hematopoietic stem cells from patients with PNH into SCID mice, they came to a different conclusion [50]. Enriched CD34(⫹) bone marrow cells from PNH patients or normal individuals were inoculated into sublethally irradiated SCID mice. The mice were supplemented with human IL-3 and stem cell factor three times weekly and human hematopoiesis was monitored. Up to 5 months, human blood cells were not detected in either group. After 7 months, human blood cells were found only in the mice that had received cells from PNH patients but not in mice bearing cells from normals. Therefore, CD34(⫹) cells from PNH patients appeared to have a growth advantage over normal CD34(⫹) cells in SCID mice. Most of the transplanted CD34(⫹) cells from PNH patients were GPI deficient, and it was therefore not possible to compare the hematopoietic activities of GPI-deficient and -sufficient CD34(⫹) cells from the same patient. The authors concluded that PNH cells have an intrinsic growth advantage rather than a survival advantage against some suppressive mechanism, such as immunologic attack. Because SCID mice have natural killer cells, an alternative explanation would be that GPI-


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negative human cells are more resistant than normal cells to mouse natural killer cells, which would be a survival advantage. The use as hosts of NOD-SCID mice, which are deficient in natural killer cells [51] would clarify this concern.

Conclusions and Perspectives The results from in vitro hematopoietic differentiation experiments with PIGA(⫺) ES cells and from chimeric mice generated using PIGA(⫺) ES cells suggest that PIGA(⫺) HSC do not have an abnormal intrinsic ability to clonally expand. These data support the hypothesis that a second factor is involved in the clonal dominance seen in patients with PNH. One possibility is that this second factor also resides within the PIGA(⫺) HSC. Alternatively, selection would be mediated by the environment. A second intrinsic factor could be an additional genetic change(s) in the PIGA(⫺) HSC. Expanded PIGA(⫺) clonal cells may be analogous to cells of a benign tumor, bearing some other gene abnormality conferring a growth advantage or increased resistance to apoptosis. If a candidate gene abnormality is found in patient’s cells, a mouse model would be useful to test its functional effect on HSC in vivo, as mice bearing such gene abnormality plus a PIGA mutation can be generated. Regarding the second possibility, the additional factor could be an abnormal condition under which the PIGA(⫺) HSC has a survival advantage over normal cells (52): autoimmunity to the HSC would be one candidate [53]. If autoantibodies against specific GPI-anchored proteins expressed on HSC are generated, normal HSC but not PIGA(⫺) HSC would be recognized and targeted. Alternatively, if cytotoxic T cells that recognize some antigen presented on the HSC are active, PIGA(⫺) HSC might be less sensitive to lymphocyte-mediated killing due to lack of a putative GPI-anchored adhesion molecule involved in the target–effector association. Mouse models should be useful to test whether autoimmune mechanisms can selectively damage normal HSC resulting in survival of PIGA(⫺) HSC.

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6. Miyata, T., Takeda, J., Iida, Y., Yamada, N., Inoue, N., Takahashi, M., Maeda, K., Kitani, T., and Kinoshita, T. (1993). Cloning of PIG-A, a component in the early step of GPI-anchor biosynthesis. Science 259, 1318–1320. 7. Stevens, V. L., and Raetz, C. R. (1991). Defective glycosyl phosphatidylinositol biosynthesis in extracts of three Thy-1 negative lymphoma cell mutants. J. Biol. Chem. 266, 10039–10042. 8. Sugiyama, E., DeGasperi, R., Urakaze, M., Chang, H. M., Thomas, L. J., Hyman, R., Warren, C. D., and Yeh, E. T. H. (1991). Identification of defects in glycosylphosphatidylinositol anchor biosynthesis in the Thy-1 expression mutants. J. Biol. Chem. 266, 12119–12122. 9. Iida, Y., Takeda, J., Miyata, T., Inoue, N., Nishimura, J., Kitani, T., Maeda, K., and Kinoshita, T. (1994). Characterization of genomic PIG-A gene: A gene for GPI-anchor biosynthesis and paroxysmal nocturnal hemoglobinuria. Blood 83, 3126–3131. 10. Bessler, M., Hillmen, P., Longo, L., Luzzatto, L., and Mason, P. J. (1994). Genomic organization of the X-linked gene (PIG-A) that is mutated in paroxysmal nocturnal haemoglobinuria and of a related autosomal pseudogene mapped to 12q21. Hum. Mol. Genet. 3, 751–757. 11. Takeda, J., Miyata, T., Kawagoe, K., Iida, Y., Endo, Y., Fujita, T., Takahashi, M., Kitani, T., and Kinoshita, T. (1993). Deficiency of the GPI anchor caused by a somatic mutation of the PIG-A gene in paroxysmal nocturnal hemoglobinuria. Cell 73, 703–711. 12. Kawagoe, K., Kitamura, D., Okabe, M., Taniuchi, I., Ikawa, M., Watanabe, T., Kinoshita, T., and Takeda, J. (1996). GPI-anchor deficient mice: Implications for clonal dominance of mutant cells in paroxysmal nocturnal hemoglobinuria. Blood 87, 3600–3606. 13. Dunn, D. E., Yu, J., Nagarajan, S., Devetten, M., Weichold, F. F., Medof, M. E., and Young, N. S. (1996). A knock-out model of paroxysmal nocturnal hemoglobinuria: Pig-a- hematopoiesis is reconstituted following intercellular transfer of GPI-anchored proteins. Proc. Natl. Acad. Sci. USA 93, 7938–7943. 14. Rosti, V., Tremmi, G., Soares, V., Pandolfi, P. P., Luzzatto, L., and Bessler, M. (1997). Murine embryonic stem cells without pig-a gene activity are competent for hematopoiesis with the PNH phenotype but not for clonal expansion. J. Clin. Invest. 100, 1028–1036. 15. Bessler, M., Mason, P. J., Hillmen, P., Miyata, T., Yamada, N., Takeda, J., Luzzatto, L., and Kinoshita, T. (1994). Paroxysmal nocturnal haemoglobinuria (PNH) is caused by somatic mutations in the PIG-A gene. Embo. J. 13, 110–117. 16. Dacie, J. V., and Lewis, S. M. (1972). Paroxysmal nocturnal haemoglobinuria: Clinical manifestations, haematology and nature of disease. Ser. Haematol. 3, 3–23. 17. Rotoli, B. R., and Luzzatto, L. (1982). Decreased number of circulating BFU-Es in paroxysmal nocturnal hemoglobinuria. Blood 60, 157–159. 18. Dessypris, E. N., Clark, D. A., McKee, L. C. J., and Krantz, S. B. (1983). Increased sensitivity to complement or erythroid and myeloid progenitors in paroxysmal nocturnal hemoglobinuria. N. Engl. J. Med. 309, 690–693. 19. Maciejewski, J. P., Sloand, E. M., Sato, T., Anderson, S., and Young, N. S. (1997). Impaired hematopoiesis in paroxysmal nocturnal hemoglobinuria/aplastic anemia is not associated with a selective proliferative defect in the glycosylphosphatidylinositol-anchored protein-deficient clone. Blood. 89, 1173–1181. 20. Robb, L. (1997). Hematopoiesis: origin pinned down at last?, Curr. Biol. 7, R10–R12. 21. Dzierzak, E., Sanchez, M. J., Muller, A., Miles, C., Holmes, A., Tidcombe, H., and Medvinsky, A. (1997). Hematopoietic stem cells: Embryonic beginnings. J. Cell Physiol. 173, 216–218. 22. Moore, M. A. S., and Metcalf, D. (1970). Ontogeny of the haemopoietic system: Yolk sac origin of in vivo and in vitro colony forming cells in the developing embryo. Br. J. Haematol. 18, 279–296. 23. Godin, I. E., Garcia-Porrero, J. A., Coutinho, A., and Dieterlen-Lievre, F. (1993). Para-aortic splanchnopleura from early mouse embryos contains B1a cell progenitors. Nature 364, 67–70. 24. Medvinsky, A. L., Samoylina, N. L., Muller, A. M., and Dzierzak, E. A. (1993). An early preliver intraembryonic source of CFU-S in the developing mouse. Nature 364, 64–67. 25. Dzierzak, E., and Medvinsky, A. (1995). Mouse embryonic hematopoiesis. Trends Genet. 11, 359–366. 26. Godin, I., Dieterlen-Lievre, F., and Cumano, A. (1995). Emergence of multipotent hemopoietic cells in the yolk sac and paraaortic splanchnopleura in mouse embryos, beginning at 8.5 days postcoitus. Proc. Natl. Acad. Sci. USA 92, 773–777. 27. Medvinsky, A., and Dzierzak, E. (1996). Definitive hematopoiesis is autonomously initiated by the AGM region. Cell 86, 897–906.


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7 The Function of GPI-Anchored Proteins Ian Okazaki and Joel Moss Pulmonary-Critical Care Medicine Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892-1434

Introduction The proteins linked to the plasma membrane by a glycosylphosphatidylinositol (GPI) anchor (Fig. 7-1) have many roles. Some possess enzymatic activity, some are involved in specific protein–protein (cell–cell, cell–matrix) interactions, and some serve as differentiation markers or as membrane receptors [1, 2]. The proposed biological functions of GPI anchors are similarly diverse, but some specific functions have been recognized (Table 7-1). GPI-linked proteins can be released from the cell surface by phospholipases C and D, to generate the signaling molecules diacylglycerol and phosphatidic acid [3, 4]. Some of the effects of insulin may be mediated in this manner. Several investigators have demonstrated the clustering of GPI-linked proteins following reaction with specific antibodies and cross-linking anti-immunoglobulin antibodies. GPI-anchor clustering increased tyrosine phosphorylation and cytokine production and induced T cell proliferation, among other effects [5, 6]. In some proteins (e.g., Thy-1, Ly-6, Qa-2), the GPI anchor was critical for signal transduction, because a protein generated by fusing the extracellular domain of the GPI-linked protein with the transmembrane domain of the class I major histocompatibility complex (MHC) H2-Db antigen failed to function in its normal signaling pathway [7]. Antibody-mediated clustering of GPI-anchored proteins resulted in their association with 60-nm membrane invaginations known as caveolae [8]. Localization of members of the src-family of non-receptor-associated protein tyrosine kinases (PTK) in caveolae is consistent with the detection of these kinases in immunoprecipitates of GPI-linked proteins. PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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Figure 7-1 Structure of GPI-anchored proteins. The core of the GPI anchor is composed of phosphoethanolamine linked to mannose residues (oligosaccharide), which in turn are attached to glucosamine (GlcN). The protein is attached to the outer leaflet of the plasma membrane through phosphatidylinositol. GPI-anchored proteins are released from the membrane by the action of glycosylphosphatidylinositol-specific phospholipase D (GPI-PLD) or phosphatidylinositol-specific phospholipase C (PI-PLC).

Following synthesis, GPI-anchored proteins associate with microdomains or ‘‘rafts,’’ which are enriched in glycosphingolipids and cholesterol and are insoluble in nonionic detergents [9]. Raft formation appears to begin in the Golgi and serves as a mechanism for sorting the GPI-linked molecules for transport to the apical surface of the cell. Once exported, the protein is anchored to the outer

TABLE 7-1 Known Functions of GPI-Anchored Molecules Mammalian enzymes Acetylcholinesterase Alkaline phosphatase CD73 (5⬘-nucleotidase) Mammalian dipeptidase Lipoprotein lipase ART1 ART2 (RT6) Leukocyte antigens Thy-1 Ly-6 (TAP) Qa-2 Sca-2 ART1

ART2 (RT6) CD14 CD24 CD48 (sgp-60) CD52 (CAMPATH-1) CD55 (decay-accelerating factor) CD59 CD73 (5⬘-nucleotidase) CD87 (uPAR) Neural cell antigens NCAM F3/F11 TAG-1 BIG-1

CNFTFR움 Cerebroglycan Ceruloplasmin Receptors/adhesion molecules Folate-binding protein NCAM F3/F11 TAG-1 BIG-1 CNFTFR움 Glypicans Cerebroglycan CD14 CD87 (uPAR) LFA-3

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leaflet of the plasma membrane where it potentially has a high lateral mobility relative to transmembrane proteins [10]. Lateral mobility may be important for positioning GPI-anchored proteins in close proximity to other molecules, such as non-receptor-associated PTK involved in specific signaling pathways. The GPI-anchored axonal glycoprotein TAG-1 promotes adhesion and extension of neurites from embryonic DPG neurons [11]. Another family of GPI-linked proteins involved in adhesion is the glypicans, which are members of the heparin sulfate proteoglycan family [12]. The folate-binding protein and the urokinasetype plasminogen activator receptor (uPAR) are examples of GPI-linked receptor molecules. The membrane-associated folate-binding protein is required for the cellular uptake of folic acid and other related metabolites [13, 14]. uPAR binds and activates urokinase-type plasminogen activator and also modulates integrin binding to fibronectin [15, 16]. Based on these data, uPAR has been proposed to play a role in fibrinolysis or tissue remodeling as well as in cell migration and tumor metastasis. In this chapter we discuss the role of GPI-anchored proteins in cell function in further detail.

GPI-Linked Proteins and Cell Signaling Role of GPI-Anchored Proteins in Lymphocyte Activation Some of the effects of GPI-anchored proteins on lymphocytes were initially observed in experiments on the murine Ly-6 family of alloantigens, which are expressed on T and B cells, thymocytes, macrophages, neutrophils, and bone marrow cells. Antibody cross-linking of Ly-6.2A (TAP), a major Ly-6-encoded protein on T cells, stimulated T cell proliferation and interleukin (IL)-2 secretion and increased intracellular Ca2⫹ accumulation [17]. Moreover, this effect was synergistic with that of T cell receptor (TCR) stimulated by anti-TCR antibody. The response to cross-linking of Ly-6.2A was absent in T cell hybridomas that did not express Ly-6.2A. These mutant T cell clones were also deficient in TCRinduced activation. Conversely, T cell hybridomas lacking TCR/CD3 complex, but not Ly-6.2A, were unresponsive to stimulation with anti-Ly-6.2A monoclonal antibody, consistent with the hypothesis that the Ly-6 and TCR pathways of T cell activation are functionally linked. The TCR/CD3-negative clones generated IL-2 in response to phorbol ester (PMA), an activator of protein kinase C, and to calcium ionophore, suggesting that the functional defect in TCR/CD3 mutants was proximal to generation of stimulatory intracellular receptor signals [18]. Antisense oligonucleotides directed against Ly-6A mRNA inhibited T cell proliferation in response to concanavalin A (Con A), anti-Ly-6A, or anti-CD3 monoclonal antibodies, but not to the combination of PMA and ionomycin [19]. Likewise, Ly-6A antisense oligonucleotides inhibited proliferation of an MHCrestricted T cell clone in the presence of antigen and antigen-presenting cell. Despite an increase in IL-2 production following cross-linking of Ly-6 antigen, Codias et al. demonstrated that costimulation of Ly-6 and the TCR/CD3 complex led to inhibition of IL-2 production in T cell hybridomas and peripheral T cells [20, 21]. Incubation of T cell hybridomas with soluble anti-CD3 and antiLy-6 or incubation of peripheral T cells with alloantigen and cross linking anti-

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Ly-6 antibodies inhibited IL-2 production. Further, inhibition of IL-2 release was not observed when T cell hybridomas were incubated with alloantigen or immobilized (i.e., insoluble) anti-CD3, demonstrating a difference in responses of hybridomas and peripheral T cells. Stimulation of IL-2 production by T cells through Ly-6 antigen was dependent on expression of the ␨ chain of the TCR, whereas Ly-6-mediated inhibition was TCR-␨ independent, thus distinguishing pathways of activation and inhibition of IL-2 production by Ly-6 [21]. The significance of the GPI anchor was demonstrated by converting Ly-6E antigen into a transmembrane protein in which the hydrophobic signal sequence was replaced with the transmembrane and cytoplasmic domains of the MHC class I Db protein. The transmembrane Ly-6EDb fusion protein failed to mediate T cell proliferation and IL-2 production [22]. When Ly-6-negative murine EL4J cells transfected with the chimeric construct were stimulated with anti-CD3 and anti-Ly-6E monoclonal antibodies, however, IL-2 production was blocked, demonstrating that, unlike the direct induction of IL-2 by Ly-6 stimulation, the prevention of anti-CD3-mediated IL-2 production by Ly-6E does not require the GPI anchor [23]. Ly-6-mediated inhibition of IL-2 production was also observed in Ly-6-transformed Jurkat cells, consistent with the fact that the machinery for Ly-6 signal transduction is also present in human cells. Antibodies to thymic shared antigen (TSA-1) or stem cell antigen-2 (Sca-2), a member of the Ly-6 multigene family [24, 25], inhibited anti-CD3 plus accessory cell-induced IL-2 production [26]. A transmembrane form of TSA-1, similar to the Ly-6EDb construct [22], was able to inhibit IL-2 production, confirming previous findings that IL-2 inhibition was not dependent on the GPI anchor. In other experiments, tyrosine phosphorylation of the TCR ␨ chain was markedly reduced following stimulation of T cell hybridomas with anti-CD3␧ and antiTSA-1 antibodies in the presence of accessory cells. It was also reported that binding activities of transcription factors NF-␬B, AP-1, and NF-AT were reduced following T cell stimulation with anti-CD3␧ and anti-Ly-6 A/E [27]. These experiments demonstrate that proteins of the Ly-6 family are capable of mediating multiple signaling events in lymphocytes. Antibody-mediated cross-linking of other GPI-anchored T cell proteins, including Qa-2, a class I MHC antigen [5], and Thy-1, a murine T cell alloantigen [6], stimulated T cell proliferation and IL-2 production. In both cases, submitogenic concentrations of PMA were required. Transgenic mice expressing a Qa-2 antigen with the transmembrane and cytoplasmic domains of H-2Db were unable to proliferate in response to antibody stimulation [7]. Further, T cells from transgenic mice expressing a chimeric GPI-anchored form of H-2Db were stimulated to proliferate by anti-H-2Db antibodies, whereas T cells possessing the wild-type transmembrane H-2Db antigen did not. The data show that Qa-2and Thy-1-mediated T cell proliferation and IL-2 production were specifically dependent on the presence of the GPI-anchor. Immunoprecipitation of Thy-1 from hybridoma T cells and thymocytes demonstrated an association between Thy-1 and the protein tyrosine kinase p60fyn [28]. p60fyn is also associated with the TCR, and antibody cross-linking of TCR or Thy-1 led to the tyrosine phosphorylation of similar sets of proteins [29, 30]. Further, thymocytes from transgenic mice overexpressing p60fyn were more sensitive than those from wild-type mice to stimulation by anti-TCR or antiThy-1 antibodies [31].


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CD48 or sgp-60 is a GPI-anchored lymphocyte antigen that functions as the CD2 ligand [32]. A monoclonal anti-sgp-60 antibody specifically inhibited T cell proliferation, IL-2 production, and IL-2 receptor (IL-2R) expression in response to T cell activation induced by Con A plus anti-CD3 monoclonal antibody or Con A plus PMA [33]. It was subsequently shown that anti-sgp-60-mediated inhibition of T cell activation appeared to interfere with the generation of inositol 1,4,5-trisphosphate and with the release of Ca2⫹ from intracellular stores [34]. Cross-linking of sgp-60 molecules in the presence of PMA, on the other hand, resulted in marked T cell proliferation [33]. The stimulatory or inhibitory effect of anti-sgp-60 depends on the experimental conditions used, and the in vivo effects of CD48 stimulation have yet to be determined. CD73 or 5⬘-nucleotidase is a GPI-linked enzyme that functions in the purine salvage pathway. It is believed to be involved in immune function, because patients with common variable immune deficiency and congenital agammaglobulinemia have deficiencies of 5⬘-nucleotidase activity [35]. Incubation of T cells with anti-CD73 monoclonal antibodies immobilized on plastic in the presence of submitogenic concentrations of PMA or immobilized anti-CD3 antibody resulted in proliferation, IL-2 production, and IL-2R expression [36, 37]. Likewise, immobilized anti-CD73 antibody enhanced proliferation induced by soluble antiCD2. The effect of costimulation with CD3 or CD2 and CD73 was similar to that observed with CD3 and CD28 (CD28 is a well-characterized accessory molecule in T cell activation) [37, 38]. CD73 increased intracellular Ca2⫹ concentration and acted as an agonist in CD2- or CD3-mediated increases in intracellular Ca2⫹. In Jurkat cells transformed with CD73 cDNA, IL-2 production was stimulated in the presence of PMA by soluble anti-CD73 or immobilized anti-CD3 [39]. IL-2 was still produced by Jurkat cells expressing a transmembrane form of CD73, in which the extracellular portion of CD73 was fused to the transmembrane portion of human tissue factor, demonstrating that the GPI anchor may not be critical for activation through CD73. Jurkat mutants that lacked a functional TCR, p56lck, or CD45 tyrosine phosphatase, but were expressing GPIlinked CD73, did not secrete IL-2 in response to anti-CD73 plus PMA, demonstrating a linkage between TCR/CD3- and CD73-mediated T cell activation [39]. The GPI-linked decay-accelerating factor CD55 binds membrane-associated C4b and C3b and protects cells from lysis by autologous complement [40]. Crosslinking of CD55 using anti-CD55 antibodies in the presence of PMA increased T cell proliferation [41]. CD59, a GPI-linked protein that is structurally related to the murine Ly-6 family of antigens, is another membrane protein that inhibits complement-mediated lysis of erythrocytes and lymphocytes [42]. An anti-CD59 monoclonal antibody in the presence of PMA and cross-linking anti-Ig caused an increase in intracellular Ca2⫹ concentration, inositol phosphate turnover, and IL-2 production in Jurkat cells. IL-2 expression required an intact TCR/CD3 complex [43]. When CD59-deficient promonocytic U937 cells were incubated with purified CD59 from erythrocytes, CD59 clustering and CD59-PTK complex formation were observed in a time-dependent fashion [44]. Further, antibodyinduced cross-linking of CD59 led to intracellular Ca2⫹ mobilization only after cluster formation had occurred. CD24 is a 42-kDa, GPI-anchored differentiation marker on B cells and granulocytes [45, 46]. Incubation of lymphocytes in the presence of PMA and cross-linking anti-CD24 antibodies resulted in an increase in intracellular Ca2⫹

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concentration, whereas CD24 cross-linking in granulocytes resulted in a respiratory burst, as evidenced by H2O2 production [47]. Signal transduction via CD24 appears to have cell-specific consequences. A GPI-anchored 42-kDa protein on the surface of rat natural killer (NK) cells was induced following incubation with IL-2. This antigen is also expressed on rat NK-like leukemia (RNK-16) cells [48]. Reaction of anti-gp42 antibodies with RNK-16-, but not IL-2-activated NK cells, resulted in increased inositol phosphate turnover and intracellular Ca2⫹ concentration. It was hypothesized that NK and RNK-16 cells stimulated by IL-2 may utilize different pathways in responding to antigen binding and gp42 cross-linking [49]. Several GPI-linked molecules have been found to be associated with nonreceptor-associated PTK activities. Indeed, PTK activities were detected in immunoprecipitates of CD55, CD59, and CD48 from human T cells, Thy-1 and Ly-6 from mouse T cells, CD24 from B cell chronic lymphocytic leukemia (B-CLL) cells, and CD14 from monocytes. Specifically, the PTK p56 lck was detected in immunoprecipitates of CD59, CD55, CD48, and Thy-1. Whereas the transmembrane polypeptides CD4 and CD8 are physically associated with p56 lck through critical cysteine residues [50] and metal ions that stabilize the interaction [51], the association of GPI-linked proteins with p56 lck is stable in the presence of alkylating or metal ion-binding reagents [52]. Antibody cross-linking of CD59, CD55, CD48, CD24, or Thy-1 induced tyrosine phosphorylation of several cellular proteins; CD24 cross-linking also increased intracellular Ca2⫹ concentration [52]. In CD55-transfected murine thymoma (EL-4) cells, p56 lck and p59 fyn were associated with CD55. The GPI anchor of CD55 was required for tyrosine phosphorylation and subsequent IL-2 production by EL-4 cells [53]. p59fyn was also required for T cell activation via Thy-1 [54]. The importance of the GPI anchor in TCR-mediated signaling was demonstrated using GPI-deficient T cell mutant cell lines. In these cells, lower amounts of the active PTK p56 lck and p59 fyn were found in immunoprecipitates of TCR. In addition, phosphorylation of the TCR ␨ chain and ZAP-70, a 70-kDa non-receptor-associated PTK that is phosphorylated and activated following TCR/CD3 engagement [55], was decreased in cells incubated with anti-CD3␧ [56]. Other examples of GPI-anchored proteins that deliver T cell activation signals include members of the ADP-ribosyltransferase (ART) family of proteins [57]. ART1 is expressed on lymphocytes and skeletal muscle cells. This enzyme transfers the ADP-ribose moiety of NAD to an acceptor protein (arginine). In muscle cells, ART1 modifies integrin 움7 [58] and T-cadherin [59]. Inhibition of ART1 activity in chick myoblasts with meta-iodobenzylguanidine prevented myoblast fusion and differentiation [60]. In murine cytotoxic T lymphocytes (CTL), ADP-ribosylation of the extracellular domain of LFA-1 inhibited intracellular Ca2⫹ mobilization and prevented homotypic cell adhesion [61]. Further, modification of a 40-kDa cell surface substrate (p40) was associated with inhibition of PTK p56lck activity resulting in decreased CTL proliferation [62]. Thus, the enzymatic activity of the GPI-anchored ART1, and not antibody binding or cross-linking of the antigen, was responsible for the induction of intracellular signaling events. RT6 (ART2) is a rodent T cell alloantigen that possesses a deduced amino acid sequence with significant identity to that of ART1 [57]. RT6⫹ T cells can


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prevent the development of diabetes in diabetes-prone BioBreeding/Worcester (DP-BB/Wor) rats [63]. Rat RT6 possesses NAD glycohydrolase and auto-ADPribosyltransferase activity [64], whereas ART1 predominantly modifies exogenous protein substrates. Cross-linking of RT6 in the presence of PMA induced T cell proliferation in response to IL-2 and IL-4 [65, 66]. Further, treatment of cells with PMA and RT6 cross-linking antibodies enhanced expression of the IL-2 receptor 움 chain, increased the amounts of p56lck and p60fyn in immunoprecipitates of RT6, and also increased levels of phosphorylation of these PTK and other RT6-associated proteins [66]. CAMPATH-1 (CD52) antigen is a small (12 amino acid), heavily glycosylated GPI-anchored protein on human lymphocytes and monocytes [67, 68]. CD52 is an important target for monoclonal antibody-induced complement-mediated lysis of T cells [69, 70]. Anti-CD52 antibodies are being utilized clinically for in vitro depletion of bone marrow T cells to prevent graft-versus-host disease after allogeneic bone marrow transplantation [71] and in vivo for treatment of bone marrow graft rejection, lymphomas, and various autoimmune diseases [72]. In the presence of PMA, cross-linking of CD52 using monoclonal antibody and anti-Ig resulted in increased T cell proliferation and release of IL-2, interferon (IFN)-웂, and tumor necrosis factor (TNF)-움. Effects of anti-CD52 and anti-CD3 antibodies on T cell proliferation were also synergistic, confirming the association between signaling events initiated by the TCR/CD3 complex and several of the GPI-anchored antigens [73]. Proteins of the CD1 family resemble the antigen-presenting MHC molecules. Murine CD1d1 controls the maturation and function of a subset (NKT) of T cells that express markers for NK and T cells [74]. Differentiation of NKT cells depends on interaction of CD1d1 with the TCR. The major ligand of CD1d1 was demonstrated to be glycosylphosphatidylinositol through the phosphatidylinositol group [75]. It was hypothesized that GPI binding to CD1d1 occurs in the endoplasmic reticulum and that GPI may help to maintain conformation of the binding groove until it is replaced by foreign antigens which have been processed in endosomes [75].

GPI-Linked Proteins in Neural Tissues Several GPI-anchored proteins that function as cell adhesion molecules have been detected in neural cells (Table 7-1). This group of proteins belongs to the immunoglobulin superfamily and includes the neural cell adhesion molecule (NCAM) [76], TAG-1 [11], a transiently expressed 135-kDa glycoprotein on developing neurons, F3/F11 expressed in the adult hypothalamoneurohypophyseal system [77–79], and the brain-derived immunoglobulin superfamily molecule-1 (BIG-1) expressed on Purkinje cells, granule cells of the dentate gyrus, and a subset of neurons in the superficial layers of the cerebral cortex [80]. Molecules such as F3/F11, TAG-1, and BIG-1 promote neurite extension from the neuronal cell body [11, 80, 81]. Other examples of GPI-anchored proteins in neural tissue include ciliary neurotrophic factor receptor 움 (CNTFR움) [82], which binds CNTF, resulting in enhanced neuronal survival [83], and ceruloplasmin, a copper-binding protein expressed in astrocytes [84]. The GPI-linked CNTFR움 exhibits significant similar-

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ity to the transmembrane IL-6 receptor 움-chain (IL-6R움). CNTF and hematopoietic cytokines IL-6 and leukemia inhibitory factor (LIF) appear to be structurally related [85]. IL-6 bound to IL-6R움 forms a complex with gp130, which activates a specific signal transduction pathway [86, 87] similar to that in CNTFR움containing neural cells. In the immortalized rat sympathoadrenal progenitor MAH cell line, CNTF binding to CNTFR움 resulted in the inhibition of cell proliferation. Engaging CNTFR움 induced phosphorylation of cell surface proteins CLIP1 and CLIP2, which are part of the CNTF receptor complex. CLIP2 may be identical to gp130 of the IL-6 receptor complex, based on the immunoreactivity of CLIP2 with gp130-specific antibodies. Phosphorylation of CLIP1 and CLIP2 led to induction of tis 1, a DNA-binding protein [86, 88]. This signaling pathway in MAH cells in response to CNTF is very similar to that involving LIF and IL-6 in hematopoietic cells [83]. Ceruloplasmin found on the surface of astrocytes is identical to the secreted isoform present in plasma [84]. The GPI-linked isoform possesses ferroxidase activity, which appears to protect the central nervous system from oxidative injury that is evident in patients with aceruloplasminemia [89].

Role of GPI-Anchored Proteins in Insulin Action Another signaling cascade involving GPI-anchored proteins is directly related to the action of insulin in specific cell systems. Physiological concentrations of insulin or the sulfonylurea glimepiride can release GPI-anchored molecules, including alkaline phosphatase [90], heparin sulfate proteoglycan [91], 5⬘-nucleotidase [92], liproprotein lipase, and Gcel, a glycolipid-anchored cAMP-binding ectoprotein [3, 4], from muscle cells and adipocytes. Release was catalyzed by a GPI-specific phospholipase C. Further processing of the GPI anchor or carboxyl terminus of the released protein occurred because the amounts of [14C]inositollabeled proteins and anti-cross-reacting determinant (CRD) immunoreactivity declined over time. Increased glucose transport in response to insulin or glimepiride is a prerequisite for activation of the GPI-specific phospholipase in adipocytes [4]. Insulin-induced release of GPI-anchored proteins and secondary processing of the glycophospholipid anchor were hypothesized as mechanisms for generating soluble mediators of insulin action such as diacyl glycerol and phosphatidic acid [4, 90, 93]. In addition, GPI molecules that are unattached to protein are present in the outer leaflet of the plasma membrane [94], where they serve as mediators of insulin action in adipocytes [95] and nerve growth factor in cochleovestibular ganglia [96].

GPI-Anchored Proteins Function as Receptors Lipopolysaccharide Receptor CD14 is the leukocyte receptor for lipopolysaccharide (LPS) [97], which is a membrane glycolipid of gram-negative bacteria that induces septic shock [98]. Binding of LPS to CD14 in a pre-B cell line resulted in tyrosine phosphorylation


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of a 38-kDa protein, NF-␬B activation, and appearance of IgM on the cell [99]. Responses to LPS occurred in cells that had been transformed with either a GPI-anchored or a transmembrane form of CD14. Ligand binding by membraneassociated CD14 is essential for LPS-induced signaling, whereas GPI attachment of CD14 to the membrane appears not to be required [100]. This is in contrast to the absence of signaling resulting from antibody-induced cross-linking of Thy-1, Qa-2, or Ly-6 antigens that lack GPI anchors.

The Urokinase-Type Plasminogen Activator Receptor The urokinase-type plasminogen activator receptor (uPAR, CD87), which binds and activates the urokinase-type plasminogen activator (uPA), is attached to the cell membrane by a GPI anchor. Activation of uPA leads to generation of the serine protease plasmin from plasminogen in the extracellular space. In addition to its role in fibrinolysis, uPAR can modulate cell adhesion and has been proposed to play a role in cell migration and tumor metastasis. uPAR was also reported to serve as a high-affinity adhesion receptor for vitronectin [101] and to inhibit 웁1-integrin-dependent adhesion to fibronectin [15]. In monocytes, uPAR is part of a large receptor complex that includes 웁2-integrins (CD11a/CD18, CD11b/ CD18) and several PTK of the src family, including p56 lyn, p60 fyn, p56/p59hck, and p59 fgr [16, 102]. Indeed, incubation of monocytes with uPA resulted in the induction of tyrosine phosphorylation and cell migration [16, 102]. In addition to stimulating proliferation and migration of cells of a human epithelial kidney tumor line (TCL-598), monoclonal antibody-induced clustering of the uPA– uPAR complex resulted in the association of gp130 and JAK kinase with uPAR leading to phosphorylation, dimerization, and binding of STAT1 to a cis-inducible element in the c-fos gene promoter [103]. Both uPAR and CNTFR움, therefore, associate with gp130, which appears to be a linker between the GPI-anchored receptor and intracellular signaling molecules. The existence of similar linker proteins for other GPI-anchored membrane proteins that mediate cellular signaling has not been detected. Activation of signaling pathways in addition to tyrosine phosphorylation has been demonstrated following uPA–uPAR interaction. Agonist occupation of uPAR in fibroblasts stimulated cell proliferation and migration along with de novo synthesis of diacyl glycerol [104]. In U937 cells, uPARinduced monocyte adhesion involved a cAMP-dependent mechanism that was independent of PTK activation [105]. In those experiments, the carboxyl-terminal lysine of uPA was critical for induction of adhesion.

The Folate-Binding Protein Folate-binding proteins (FBP) have a high affinity for folic acid and reduced folate derivatives. Soluble and membrane-bound FBP have been identified; the membrane-associated isoforms are attached to the cell surface by GPI anchors [13, 14]. GPI-anchored FBP mediates uptake of folate compounds by an energydependent endocytic mechanism. 5-Methyl tetrahydrofolate bound to the FBP is transported to an acidic vesicular compartment, where it dissociates from the FBP before translocation into the cytosol; FBP is then recycled to the membrane surface [106, 107]. In L1210 murine leukemia cells, acylation of the inositol ring

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of the GPI anchor, a common GPI modification, resulted in greater affinity of the FBP for folate compounds and increased folate uptake [108]. Because antibody cross-linking in vitro stimulates internalization of several GPI-anchored proteins [18, 109], antibody cross-linking may simulate ligand binding in vivo and internalization of GPI-anchored molecules, as occurs in folate transport. The numbers of folate receptors on certain malignant cells are often markedly elevated [110], potentially giving the cells a selective advantage, but also providing a mechanism for chemotherapeutic intervention.

Glypicans and Glypican-like Molecules Glypican and glypican-like molecules are a family of GPI-anchored heparin sulfate proteoglycans (HSPG) [12, 111–114]. The carboxyl-terminal heparinbinding domain of fibronectin was shown to bind HSPG and integrins at the cell surface and appears to play a role in cell adhesion as well as in prevention of tumor metastasis. In fact, a cell surface glypican from mouse melanoma cells was found to bind to the synthetic fibronectin peptide FN-C/H II derived from the carboxyl-terminal heparin-binding domain [115]. Glypicans from vascular endothelial cells were found to have a higher affinity for antithrombin III than for other endothelial cell HSPG, including syndecan and fibroglycan. One possibility is that the antithrombin III–glypican interaction at the endothelial cell surface must serve as a physiologic anticoagulant [116]. Glypicans and other HSPG also bind to lipoprotein lipase [117], a major contributor to the enzymatic hydrolysis of plasma triglycerides [118]. Basic fibroblast growth factor (bFGF) is a cytokine required for stimulation of bone marrow stromal cell growth [119]. A glypican is the low-affinity binding site for bFGF [120], and this interaction modulates the action of bFGF in hematopoiesis [121]. The fact that glypicans are shed from the cell surface [12] suggests that effects of the glypican–ligand interaction in cells may be regulated by glypican release. An endogenous GPI-specific phospholipase D that released the bFGF–glypican complex has been detected in bone marrow cell cultures [122].

GPI Anchors and Membrane Structure Lateral Mobility One of the proposed major advantages of GPI-anchored proteins is unhindered lateral movement in the plasma membrane that is made possible by a lipid anchor, in contrast to the much more limited mobility of proteins with transmembrane domains. The diffusion coefficient of Thy-1, as measured by fluorescence recovery after photobleaching using a rhodamine-conjugated anti-Thy-1 monoclonal antibody, was considerably larger than those of other membrane proteins [10]. The mobility of the GPI-anchored guinea pig sperm protein (PH-20), which is involved in sperm adhesion to the egg zona pellucida, varied 250-fold during the different stages of sperm maturation but was similar to Thy-1 diffusion after the acrosome reaction [123]. The variability in diffusion of GPI-anchored proteins was thought to be due to changes in the interactions of the ectodomain of PH-20 with those


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of other membrane proteins during spermatogenesis. Lateral mobility of Qa-2 labeled with antibody-coated gold particles was greater than that of the nonGPI-anchored MHC class I molecule H-2Db, although there were barriers to lateral movement of both membrane proteins [124]. Thus, GPI-linked proteins may not always have unrestricted lateral mobility.

GPI-Anchored Proteins and Caveolae GPI-linked molecules have been reported to concentrate in plasma membrane invaginations called caveolae [8]. Caveolae appear to be the sites of entry of small molecules and ions into the cell by potocytosis [8]. In addition to GPIanchored proteins, caveolae are enriched in cholesterol, caveolin, an integral membrane marker protein for caveolae, ganglioside GM1, the 1,4,5-inositol trisphosphate receptor, Ca2⫹-dependent ATPase, and members of the src-family PTK [8], as well as proteins known to mediate vesicle formation, docking, and fusion [125]. Clustering of GPI-anchored molecules in caveolae was induced by antibody cross-linking [109, 126, 127]. The physiological relevance of GPI sequestration in caveolae has been questioned, however, because in the absence of antibody-induced clustering, GPI-linked folate receptors appeared to be diffusely distributed over the plasma membrane [127]. In lymphocytes, which lack caveolae, there was little association observed between GPI-anchored Thy-1 and the glycosphingolipid GM1 except following detergent treatment, when these molecules associated in glycolipid microdomains [128]. On the other hand, when exogenous GPI-anchored CD59 was added to CD59-negative U937 cells, CD59 spontaneously clustered and associated with PTK within 2 h. Before clustering, however, CD59 remained diffusely distributed on the cell surface and Ca2⫹ signaling was not detected until CD59 complexes had formed [44]. The observed association of GPI-anchored proteins with caveolae that contain members of the src-family of PTK (p56 lck, p59 fyn, c-Yes) and the capacity for GPI-linked proteins to generate PTK-induced signaling events in response to ligand or antibody cross-linking are consistent with the existence of signal-transducing GPI-domains [129–131]. A novel method for isolating caveolae from endothelial cells demonstrated that GPI-anchored proteins are located in microdomains separate from caveolae (Fig. 7-2; see color plate). By these techniques, GPI-linked proteins were found in an annular plasmalemmal domain surrounding the caveolar invagination [132]. Previous methods for isolating caveolae that employed nonionic detergent solubilization generated Triton X-100-insoluble membrane domains, which included caveolae, linear membrane sheets, and larger more heterogeneous vesicles in which GPI microdomains and caveolae were intermixed [132]. The physical proximity of distinct GPI microdomains to caveolae [59, 132], which contain molecular machinery for signal transduction, is nonetheless still compatible with an integrated signaling cascade following activation of GPI-linked molecules.

Apical Processing of Proteins One proposed role for the GPI anchor is as a signal for transport of proteins to the apical surface of the cell [133]. Although soluble in nonionic detergent

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immediately after synthesis, GPI-anchored proteins associate with Triton X100-insoluble glycosphingolipids and cholesterol before their transit to the cell surface. The clusters of cholesterol, glycosphingolipids, and GPI-anchored proteins are formed in the Golgi membranes and are incorporated into apex-bound vesicles [131, 134]. Upon reaching the cell surface, the GPI-enriched domains are initially clustered but disperse in a few hours [135]. Although GPI-anchored proteins are often associated with caveolae, caveolin, which is required to promote caveolar assembly, was recently found not to be involved in apical sorting of GPI-anchored proteins [136]. Rather, the redistribution of GPI-anchored proteins to caveolar regions appears to be dependent on antibody- or ligandinduced cross-linking [109, 127]. There is evidence that some GPI-anchored proteins reside in ‘‘rafts’’ (Fig. 7-2), specific subcompartments rich in cholesterol and glycosphingolipids that become detergent-insoluble as they traverse the Golgi apparatus [9, 136]. The composition of glycospingolipid fatty acyl chains has been hypothesized to be critical for raft formation [137]. Of some interest is that radiolabeling studies in polarized Madin-Darby canine kidney cells demonstrated that free GPI units, synthesized in the ER but not linked to proteins, were not directed exclusively to the apex but instead accumulated relatively equally in both apical and basolateral plasma membranes [138], a distribution distinct from that of the GPI-anchored proteins that were directed exclusively to the apical surface.

Summary Although proteins attached to the plasma membrane through a GPI linkage are certainly diverse, these proteins now appear to use common signaling pathways. The association of GPI-anchored proteins with members of the src-family PTK has been established, in particular, following antibody cross-linking of T cell antigens. It is unclear whether the effects of cross-linking antibodies in vitro are analogous to ligand binding to GPI-anchored proteins in vivo, as is observed with the TCR/CD3 complex. Molecules with GPI anchors that do not span the lipid bilayer have been assumed to form a obligatory complex with transmembrane proteins to transmit extracellular signals to intracellular effectors. CNTFR움 and uPAR are associated with gp130, a presumptive linker protein. No such linker has been identified, however, for the great majority of GPI-anchored proteins. Many GPI-linked proteins are enzymes. The effects of ectoenzyme activity on cell function have often seemed independent of GPI signaling. ART1 in lymphocytes and muscle cells appears to be unique in its ability to initiate cell signaling following modification of integrins and other membrane proteins. In some cells, proteins such as uPAR, in response to engagement of the GPIanchored protein, form large membrane-associated complexes with integrins and PTK; ART1 may be utilizing a similar mechanism of signal transduction. The effects of insulin on GPI-anchored proteins and the role of free GPIs in Ca2⫹ mobilization and inositol 1,4,5-trisphosphate turnover are consistent with the use by GPI-linked molecules of multiple intracellular signaling pathways. As more proteins are found to be linked to the membrane by GPI anchors, the diversity of functions attributed to the anchor will no doubt increase.


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8 GPI in Lower Animals Louis Schofield The Walter and Eliza Hall Institute of Medical Research, Royal Melbourne Hospital, Victoria 3050, Australia

Introduction Glycosylphosphatidylinositol-linked proteins (GPI) molecules are common to all eukaryotes. Many aspects of GPI structure, biosynthesis, and biology, however, have been elucidated from the study of lower eukaryotes, particularly the parasitic protozoa. Key structural features of this class of glycolipid were first elucidated by investigation of the membrane anchor of the variant surface glycoprotein of the parasitic protozoan Trypanosoma brucei [1, 2]. Since then, a range of GPIs have been characterized from many other protozoal taxa, including Trypanosoma cruzi [3, 4], Plasmodium [5, 6], Leishmania [7–9], and Toxoplasma [10]. Protozoa represent perhaps the most ancient and diverse trunk of the eukaryotic evolutionary tree, and the presence of core structural features of GPIs in this group as well as in plants, yeast [11], fish, and numerous mammalian sources [12, 13] attests to conservation of GPIs over 600 million years of evolution. For an excellent review of GPI structure and biosynthesis in the parasitic protozoa, see McConville and Ferguson [14]. There are, of course, several other reasons for the major role played by the study of parasitic organisms in the elucidation of GPI biology. The parasitic protozoa are unusual in that they express very high levels of GPI-anchored proteins or GPI-anchored glycoconjugates. Compared with mammalian cells, which may contain approximately 105 GPI copies per cell and for which GPIanchored proteins are a minority of cell surface proteins, various parasite species may contain up to 2 ⫻ 107 GPI copies per cell, and GPI-anchored structures clearly predominate on the cell surface. Across taxa, the majority of surface PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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proteins in the parasitic protozoa are anchored into the membrane by GPI. Examples are the membrane-form variant surface glycoprotein (mfVSG), transferrin-binding protein, and GPI-anchored trans-sialidase of T. brucei; the Ssp-4 and 1G7 major surface glycoproteins, 35/50 kDa sialic acid acceptor, and 160 kDa flagellar antigen of T. cruzi; the majority of malaria sporozoite and merozoite surface proteins, including the circumsporozoite (CS) protein, MSP-1 protease, MSP-2, and MSP-4 of Plasmodium falciparum; the gp63 metalloprotease and PSA-2 of Leishmania; and most surface antigens of Toxoplasma gondii. This represents an enormous diversity of protein structure and function, including coat proteins involved in immune evasion, receptors/ligands, and suface hydrolases. As such, there is little in common between these molecules other than the mode of membrane attachment. Nonetheless, all surface proteins of parasitic organisms are of particular interest because of their central role in the host/ parasite interaction and in the response to host immunity, and many of the proteins listed above are under consideration as vaccine candidates. For these reasons, an understanding of GPI biology in these organisms bears on the understanding and prevention of several serious human diseases. In addition to GPI-anchored proteins, parasites such as T. cruzi and Leishmania spp. also elaborate GPI-anchored nonprotein glycoconjugates such as lipophosphoglycan (LPG) and lipopeptidophosphoglycan (LPPG), and a range of GPIs and GPI-like structures which are not anchored to other structures but exist free in the plasma membrane. In Leishmania, where they are particularly abundant, these structures are referred to as glycoinositolphospholipids, or GIPLs [15]. GIPLs are present in high copy number (approximately 107 /cell) and may cover up to 80% of the amastigote parasite surface. Indeed, these molecules are also exported into the macrophage host cell [14]. In addition to being of interest in their own right, these structures have proven very informative as model systems for the elucidation of GPI biosynthetic pathways [16]. Finally, one further reason for the study of GPI biology in the parasitic protozoa is that GPIs and related structures play central functional roles as pathogenicity factors and as regulators of host cell and tissue responses. Relevant to the role of GPI anchors in all eukaryotic biology is the capacity of parasite GPIs to regulate host cell function, validating the concept of GPI-mediated signal transduction in mammalian systems. Accordingly, the bioactivity of protozoal GPIs constitutes the main focus of this chapter.

Structures of Protozoal GPIs Structurally related to phosphatidylinositol (PI), the canonical second-messenger system, protein GPI anchors from both protozoal and mammalian sources contain a conserved core glycan of the sequence Man움1-2Man움1-6Man움1-4GlcNH2, which is linked to the 6-position of the myo-inositol ring of PI. GPIs are constructed on the cytoplasmic face of the endoplasmic reticulum membrane by the sequential addition of sugar residues to PI by the action of glycosyltransferases [17]. The maturing GPI is then translocated to the luminal side of the endoplasmic reticulum [18, 19], where it may be covalently attached to proteins for export to the cell surface. The tetrasaccharide core glycan may be further substituted

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with the addition of other sugars (galactose, N-acetylglucosamine, sialic acid), phosphates, and ethanolamine groups in a species- and tissue-specific manner. In addition, there is a certain degree of structural diversity in the type and composition of the fatty acid moieties in various GPIs, which include diacylglycerols, alkylacylglycerols, monoalkylglycerols, and ceramides. Several GPIs also carry additional palmitoylations or myristoylations to the inositol ring, which renders the GPI-anchor resistant to hydrolysis by PI-specific phospholipase C. The structure of T. brucei 118 (MITat 1.5) is one of the simplest described to date, consisting of the evolutionarily conserved core glycan and a dimyristoylglycerol: ethanolamine–phosphate–Man움1,2Man움1,6Man움1,4GlcN– phosphatidyl-myo-inositol [20]. The biosynthetic pathway of the malarial GPI has now been elucidated [5, 6, 21, 22]; the pathway leads to the formation of two potential GPI membrane anchor precursors, Pfgl움 (ethanolamine– phosphate – (Man움1,2)-Man움1,2Man움1,6Man움1,4GlcN – phosphatidyl-(acyl)myo-inositol and Pfgl웁 (ethanolamine-phosphate-Man움1,2Man움1,6Man움1, 4GlcN-phosphatidyl-(acyl)-myo-inositol) [5]. Structural analysis of the core glycans of the GPI anchors of the merozoite surface protein 1 (MSP-1) and 2 (MSP-2) of P. falciparum revealed that both anchors carry a mannose in an 움1,2 linkage to the third mannose of the trimannosyl core glycan [6], indicating that Pfgl움 is their sole precursor. The hydrophobic fragment of the MSP-1 and MSP-2 GPI anchors is composed of an acylated inositol and membraneanchoring fragments consisting of diacylglycerols [6]. Compositional analysis of the radiolabeled fatty acids shows that the diacylglycerol fragments of both protein-bound GPI anchors preferentially contain [3H]palmitic acid, whereas the acylated inositol glycan fragments preferentially contain [3H]myristic acid [6]. To date, myristoylation of the inositol ring is a feature unique to malarial GPIs. When the range of GPI structures is expanded to include GIPLs of Leishmania, the conserved structural motif Man움1-4GlcNa1-6PI can still be found. GIPLs can be classified into three types depending upon their glycan structure, and they are expressed at different levels in various developmental stages: Type 1 GIPLs are essentially similar to GPI protein anchors; type 2 GIPLs are related in having the core sequence Man움1-4GlcN1-6myo-inositol but diverge thereafter in linkage to a Galf웁1–3Man움1-3 structure (unusual among eukaryotic GPIs in modification of the otherwise evolutionarily conserved core glycan structure). Unlike protein anchor GPIs, the type 2 GIPL sequence forms the basis of the LPG membrane anchor. Finally, hybrid GIPLs have the conserved glycan sequence but contain an additional mannose residue in an 움1-3 linkage to the proximal mannose [14]. As we will discuss below, an important feature of the GPI structures elaborated by Leishmania and T. cruzi, in contrast to those of P. falciparum and T. brucei, is the preponderance of alkylacylglycerols, lyso-alkylglycerols, and ceramides, as opposed to diacylglycerols, within the lipid domain. The overall picture concerning GPIs is of a clearly related family of glycolipids sharing certain evolutionarily conserved core features but with a high level of variation in fatty acid composition and side-chain modifications to the glycan. In the case of type 2 GIPLs of Leishmania, even the otherwise evolutionarily conserved core glycan has been modified to accept the highly

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unusual Gal-furanose modification. The only feature that consistently distinguishes protozoal from metozoan GPIs is that the latter are substituted with additional ethanolamine phosphates. The functional significance of the structural differences among GPIs is not yet clear, but as we have demonstrated elsewhere [23–25] and will discuss below, there is evidence that these differences among GPIs impart characteristic properties of signal transduction to this class of glycolipid.

Evidence for GPI-Mediated Signaling in Mammalian Cell Biology Signaling Mediated by GPI-Anchored Proteins Originally, GPIs were thought to function solely as a novel type of membrane anchor for proteins. However, recent studies in mammalian cell biology have implicated GPIs as mediating signal transduction within cells. Ligation, clustering, or antibody-mediated cross-linking of a range of GPI-anchored proteins with diverse functions in lymphoid and myeloid biology, such as GPI-anchored Thy1 [26–28], Qa-2 [29, 30], CD24 [31], Ly-6/TAP [32], CD59 [33, 34], Sgp-60 (CD48) [35], and CD73 [36, 37], imparts transmembrane regulatory signals to T lymphocytes [38], leading to various downstream responses, including proliferation, cytokine expression, and T cell anergy. In macrophages, cross-linking GPIanchored proteins such as CD14 [the lipopolysaccharide (LPS) receptor] and CD16 (Fc-웂RIIIB) induce macrophage activation, leading to cytokine expression and the oxidative burst [39, 40]; nonreceptor protein tyrosine kinases (PTKs) are implicated in this phenomenon. Cross-linking of certain GPI-anchored proteins induces rapid onset tyrosylphosphorylation of intracellular substrates, and coprecipitation experiments show that various GPI-anchored proteins are physically associated with the src-family kinases fyn, lck [41–43] fgr, lyn, and hck [44]. Furthermore, when certain proteins that are normally GPI-anchored are expressed with artificial transmembrane domains, the capacity to activate cells following perturbation is lost [29], indicating a requirement for the GPI anchor in these phenomena; in neutrophils, perturbation of Fc-웂RIIa together with GPIanchored Fc-웂RIIIB or unrelated GPI-anchored proteins enhances calcium flux in a manner that depends upon the GPI anchor [45]. Nonetheless, this body of data provides only circumstantial evidence that GPI molecules are cell signal mediators. As outlined above, most observations implicating GPIs in signal transduction are indirect, based on the experimental cross-linking of GPI-anchored proteins with antibodies, often in the presence of phorbol esters [46]. The physiological relevance of this type of perturbation is doubtful, and the signaling activity observed in T lymphocytes under these conditions may reflect an artifactual disruption of the T cell receptor (TCR) signaling complex. Furthermore, a topological objection remains: because the cellular signaling machinery is located in the cytosol and GPI-anchored proteins are located in the outer (luminal) leaflet of the plasma membrane, how a GPImediated activation signal is transmitted across the membrane to the interior of the cell remains unclear. The contribution of GPI moieties to cell signaling may


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merely reflect the increased lateral mobility of GPI-anchored proteins in the plasma membrane compared to that of transmembrane forms of the molecule, allowing the formation of a macromolecular complex in which signaling is effected by other molecules. In some [47] but not all [29, 45] cases, the use of transfection methods to express molecules with either GPI anchors or transmembrane domains has shown that GPIs are not required for the mediation of biological responses to ligand. One plausible interpretation of these studies (and a model of signaling mediated by GPI-anchored proteins) is that these molecules bind in cis to transmembrane partners, which themselves effect intracellular signaling, and that the specificity is imparted by the extracellular domain of the GPIanchored protein together with its ligand. The contribution of the GPI moiety may be as a membrane localization device or to facilitate lateral movement and clustering. For the ciliary neurotrophic factor receptor [48] and glial cell neurotrophic factor receptor [49], a complex forms by association of ligand with a GPI-anchored protein that binds in cis to transmembrane signal-transducing partners. The GPI anchor is not required for this activity, as signaling can occur when the anchor is replaced by a transmembrane domain or even by soluble protein. Nonetheless, these considerations do not rule out a model of GPImediated signaling in which the GPI anchor itself binds to an unidentified linker protein and/or is hydrolyzed by phospholipases to generate lipidic second messengers. Indeed, both types of activity may occur, and GPIs may provide unsuspected ancillary or accessory signals to more conventional pathways. One likely consequence is that within the cell there would be relative lack of specificity provided by GPI signaling. This may be the case in cells of hematopoietic origin such as T lymphocytes, where clustering of a wide variety of different GPIanchored proteins leads to cell activation. Taken together, therefore, studies in mammalian systems that have sought to investigate GPI function by perturbation of GPI-anchored proteins are not to distinguish satisfactorily able between effects resulting from a contribution by the protein component, the purely physical properties of the GPI moieties (lateral diffusion), other signaling components, and those that may be due to GPI-mediated signaling itself.

GPI-Derived Mediators of Hormone Action? In addition to the evidence obtained from GPI-anchored proteins, a considerable body of literature suggests a role for GPI molecules as precursors for second messengers in postreceptor signaling in a variety of hormone and cytokine systems. The specific binding to receptors of agents such as insulin [50–52], nerve growth factor [53, 54], transforming growth factor 웁 [55], interleukin-2 [56, 57], thyrotropin [58], follicle-stimulating hormone, and chorionic gonadotropin [59] reportedly leads to the hydrolysis of molecules with GPI-type features and the release of pharmacologically active GPI hydrolysis products such as inositolphosphoglycan (IPG). As these putative IPG moieties are able to mimic some of the bioactivities of hormones and cytokines, GPI hydrolysis products may serve as a novel class of extracellular second messengers or to mediate signal transduction in response to hormones and cytokines. Further evidence in support of a role for GPI molecules in postreceptor insulin action came from the use of antisera to the myo-inositol 1,2-cyclic phosphate (cross-reactive determinant) on GPI

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anchors, which inhibit insulin action [60]. Insulin can also be antagonized by partial GPI structures [61]. Most importantly, insulin activity was examined in glycosyltransferase mutant, GPI-negative K562 cell lines [62]. The IA mutant, which fails to make GlcNAc-PI, and the IVD mutant, which fails to de-Nacetylate GlcNAc-PI to GlcN-PI, were both profoundly impaired in glycogen synthesis, although insulin receptor autophosphorylation, IRS-1 phosphorylation, and interaction with Grb2 were unaffected [62]. There are as yet no alternative explanations for these observations within the classical framework of postreceptor insulin signal transduction. Despite this voluminous literature implicating GPI hydrolysis as a step in postreceptor signaling by hormones and cytokines, no structure of any such hormone-sensitive mammalian GPIs has yet been determined. The description of these putative second messengers as GPI-derived inositolphosphoglycans remains speculative, based mostly upon the release from cells of molecules that incorporate labeled precursors such as glucosamine, mannose, and inositol, consistent with a GPI structure. Indeed, the biological activities ascribed to IPG fragments cannot yet be assigned unambiguously to homogeneously or compositionally pure chemical moieties of any description. The proof that GPIs participate in these important biological processes still awaits the satisfactory purification and structural characterization of such molecules.

Membrane Microdomains The recent discovery of the complex biology of caveolar structures [63] or lipid rafts [64] within cell membranes raises further questions of the potential role of GPI moieties in cell signaling. In higher eukaryotes (but not in protozoa), cell membranes contain phase-separated ‘‘rafts’’ with highly unusual lipid compositions (see Chapter 10 by Mayor). Enriched in cholesterol and glycosphingolipids, these structures are also resistant to solubilization by Triton X-100 at 4⬚C and have a light buoyant density. Initially, interpretation of experimental results such as detergent insolubility was controversial and there were doubts that this property could be taken as indicative of an underlying biological entity. However, it is now accepted that specialized microdomains do exist at the cell surface, albeit of small size, and moreover that these structures probably constitute specialized signal transduction devices [65, 66]. Caveolar complexes or rafts appear highly enriched in molecules involved in signal transduction. In addition to gangliosides, sphingomyelin, ceramide, and diacylglycerol, they contain acylated proteins such as heterotrimeric G proteins; src-family protein tyrosine kinases (PTKs) (Fyn, Hck, Lyn); caveolin [67]; prenylated proteins such a Ras and Rap; receptors such as those for insulin, PDGF, and EGF; signal transducers such as PKC움, SOS, and Grb2; MAP kinases; PI3 kinase; Raf1; phosphoinositides; and membrane transporters such as IP3 receptor and both Ca2⫹ and H⫹ ATPases. Furthermore, on the luminal side, they are highly enriched in GPI-anchored proteins. Mutations that abolish acylation sites or GPI anchorage abrogate the association of proteins with these structures. Indeed, GPI anchorage and acylation are the only known signals for targeting proteins to rafts, although other signals must exist, as some raft proteins such


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as 웁-integrins [68] contain neither modification. With few exceptions, association of PTKs with rafts requires their dual myristoylation and palmitoylation. These unique structures therefore serve among other functions to compartmentalize many signaling pathways, and it is likely that they also represent areas where signaling cross-talk, regulation, and integration can occur. Although the association of GPI-anchored proteins with rafts does not resolve the question of whether GPIs represent merely localization (raft-targeting) devices or specific signal transduction agents, it is tempting to speculate that GPI-specific signaling events can occur through novel mechanisms involving the multiple signaling partners identified in rafts. On balance, therefore, and despite some caveats that must be placed on the interpretation of the literature, GPIs clearly represent a class of glycolipid with potentially great importance in the regulation of cell function through novel pathways of signal transduction. However, the evidence from mammalian systems remains equivocal. Because GPIs are elaborated so frequently by parasitic protozoa, both free and in association with surface antigens, we and others have sought to investigate a possible role for these molecules as signaling agents in the host/ parasite relationship. To this end, GPIs and GIPLs of various parasite taxa were purified and examined for their capacity to regulate host cell function [23–25, 69–73]. In contrast to authors of studies that rely upon the cross-linking of GPIanchored proteins, we were able to show that protozoal GPIs are able to activate endogenous mammalian GPI-based signal transduction pathways when added alone to cells as structurally defined signaling agonists and that these activities may account for much pathology and indeed immune dysregulation in protozoal infections. These findings provide proof of cellular bioactivity by GPIs and are discussed below, with particular reference to studies on macrophages and lymphocytes, although it should be noted that other cell types such as hepatocytes [69] and adipocytes [70] may also be influenced in diverse ways by parasite GPIs.

Signal Transducton in Host Cells by Protozoal GPIs As noted above, P. falciparum and T. brucei MITat 1.5 [1, 2] elaborate some of the simplest and most straightforward of GPI structures. In T. brucei, the GPI glycan consists of the evolutionarily conserved backbone ethanolamine– phosphate–6Man움1-2Man움1-6Man움1-4GlcN1-6myo-inositol. In Plasmodium, this is substituted with an additional terminal mannose in an 움1-2 linkage, and the GPI also bears a myristoylation to the inositol, rendering GPIs resistant to hydrolysis by PI-specific phospholipase C [5, 6]. Both GPIs also contain saturated fatty acids (dimyristoyl and dipalmitoyl, respectively) in an sn-1,2-diacylglycerol [6]. Saturated sn-1,2-diacylglycerols are the physiological activators of PKC [74]. We sought to determine whether these simple GPI structures could mediate signal transduction in host cells and to investigate the mechanism of such a process. Based on the proposed association of mammalian GPI-anchored proteins with src-family protein tyrosine kinases, we first demonstrated that GPI molecules induce the very rapid activation of tyrosine kinase activity in murine LPS-nonresponsive macrophages and human vascular endothelial cells [23, 24], leading to the rapid tyrosine phosphorylation of multiple intracellular substrates. Tyrosyl-

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phosphorylation appeared within 30 s to 1 min after the addition of low concentrations of GPI to cells (10 nM–1 애M), peaked at about 10–15 min, and disappeared after 20–30 min. We confirmed that the tyrosine kinase activity was due to nonreceptor PTKs by use of low concentrations of specific PTK antagonists (herbimycin A, tyrphostin), which were able to block phosphorylation, GPIinduced cell signaling, and downstream gene expression [23, 24]. The activation of src-family PTKs is considered a key signaling step in the regulation of numerous cellular responses, including macrophage and lymphocyte activation. The pattern of tyrosylphosphorylation substrates detected strongly suggests that the kinase activity induced by GPI results from the activation of the macrophage lineagespecific kinases p59/61hck and p56/59lyn. Immunoprecipitation and in vitro kinase assays showed that GPIs also upregulate p59/61hck kinase activity within 30 s to 1 min of addition to macrophages, as measured by autophosphorylation and transphosphorylation of an exogenous substrate—the earliest measurable event in GPI-mediated macrophage activation [25]. PTK activation appears to be mediated by binding of the evolutionarily conserved GPI core glycan to an uncharacterized glycan-specific receptor on the surface of cells. GPI-associated fatty acids and phosphate groups are not required for PTK activation, because a PLA2-generated lyso-GPI-based hydrolyzed inositolphosphoglycan and GPI-specific phospholipase D deacylated inositolglycan are each able to activate PTKs [25]. Moreover, the activation of PTKs maps to the complete glycan moiety of the GPI. Using structurally defined GPI precursors which differ in the degree of glycosylation, it has been possible to show that the minimal structural requirement for PTK activation and downstream cell activation is the evolutionarily conserved Man3-GlcN-inositol; Man2-GlcN-inositol is not active [25]. It is therefore possible that the conservation of the core glycan structure reflects structural constraints on this signaling activity. Although lyso-GPI and the inositolglycan fragment are sufficient to activate PTKs, they do not provide the full range of signaling events leading to downstream gene expression and macrophage activation, indicating that an additional non-PTK signal is necessary. The sn-1,2-diacylglycerols within the P. falciparum and T. brucei GPIs, which independently activate cellular protein kinase C (PKC), are required. Of the PKC family isoforms present in macrophages (the calciumdependent 움, 웁, 웂, the novel 웃 and ␧, and ␪), GPI activates only the calciumindependent ␧ isoform, demonstrating the highly specific nature of the response at the molecular level [25]. PKC activation is blocked by selective deacylation of the GPI and by specific PKC antagonists. The same antagonist, calphostin C, blocks TNF and iNOS expression [23, 24]. As mentioned above, deacylated GPI (lyso-GPI and inositolglycan) is insufficient to activate PKC or to induce downstream effects. Thus, malarial and trypanosomal GPIs impart at least two distinct signals to cells through structurally distinct elements (PTK activation via the glycan and PKC activation via diacylglycerol), both of which are required for the pathophysiological effects of GPI. Macrophage activation can be induced by both inositolglycan and diacylglycerol moieties of GPIs of both P. falciparum and T. brucei [25, 71, 75]. Remaining issues concerning our two-signal model for GPI-specific signaling events are the identities of the glycan-specific receptor responsible for activation of PTKs and the mechanism by which GPI-derived


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fatty acids enter the signaling pathway as agonistic or antagonistic second messengers. As malarial GPI is myristoylated on the inositol and resistant to hydrolysis by PI-PLC [5, 6, 21], a possibility is that GPI is hydrolyzed at the cell surface by GPI-specific phospholipase D, acting as a PTK-dependent signal-activated phospholipase. Alternatively, the intact GPI may be translocated from the external surface of the cell to the interior, similar to processes during GPI biosynthesis, where a ‘‘flippase’’ has been postulated to account for the movement of GPI precursors from the cytoplasmic to the luminal face of the endoplasmic reticulum [19]. It also remains to be determined whether GPIs containing ceramides are hydrolyzed to physiologically enter the sphingomyelinase pathway.

Parasite GPIs as Functional Toxins and Pathogenicity Factors GPI of P. falciparum was first shown to exert regulatory effects on host cells and act as a parasite toxin by its ability to induce TNF and IL-1 production by macrophages [70]; this activity required GPI-associated diacylglycerol and could be blocked by the PKC-specific antagonist staurosporine, indicating the initiation of GPI-derived fatty acid-dependent signaling pathway. Both P. falciparum and T. brucei GPIs in mice induce a transient pyrexia [70, 71] and can cause the death of recipients through TNF-mediated cachexia [70]. The induction of these cytokines by a parasite toxin is thought to be responsible for much malarial morbidity and mortality [76, 77]; malarial GPI is thus a major pathogenicity and virulence determinant. GPI purified from the variant surface glycoprotein of T. brucei had similar activities in macrophage activation [71] and could also account for the high level of IL-1 and TNF found in trypanosomiasis, which also mediate disease states in this infection. Malarial and trypanosomal GPIs induce a range of other activities in various tissues and cell types, which may account for several important pathophysiological syndromes and conditions in these two diseases—and also influence immune competence. Malarial GPI directly, and in synergy with interferon-웂, increases inducible nitric oxide synthase (iNOS) gene expression and NO output, implicated in the etiology of the cerebral malaria syndrome [23], and also upregulates ICAM and VCAM expression in host cells [24]. ICAM acts as a host ligand recognized by parasites in vascular sequestration and is therefore also implicated in the etiology of cerebral malaria [78]. Importantly, crude total parasite extracts, which also induce TNF expression by macrophages, activate the same signals as parasite GPIs: hck, lyn, PKC␧, and NF-kB/c-rel activation, and with identical kinetics; thus, GPIs alone appear sufficient to activate the signaling pathways leading to TNF and iNOS expression. The biological activity of total parasite extracts, including TNF output, can also be inhibited by high concentrations of GPI partial structures, such as glucosamine, mannose, and phosphatidylinositol [70, 79]. Most importantly, monoclonal antibodies to malarial GPI neutralize iNOS [23], ICAM-1 [24], and TNF [25, 80] induction by parasite extracts, clearly demonstrating that GPI is a necessary and major toxin of parasite origin and responsible for these host responses. In its

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role as a malarial toxin, GPI contributes significantly to global mortality and morbidity.

Regulation of Macrophage Function by Glycoconjugates of Leishmania Clearly, cell signaling events map to salient structural features of the malaria and T. brucei GPIs. These findings are doubly significant because GPIs from other parasitic taxa, particularly Leishmania and T. cruzi and certain yeast, show quite distinctive structural features that could be expected to have distinct signaling activities within cells. In other words, there may exist signaling specificity associated with structurally distinct GPIs from various taxa, including protozoa such as Plasmodium, Leishmania, T. brucei, T. cruzi, and Toxoplasma, but also extending to yeast and fungi. This is ancillary to the larger hypothesis that throughout eukaryotic biology various GPIs mediate a wide range of crucial regulatory events and that different pathogens have evolved diverse GPIs to regulate host cell function to their advantage. We and others [81, 82] have obtained evidence for distinct signaling activities mediated by GPIs of various parasite species. As mentioned above, Leishmania contain various structurally bizarre glycoinositolphospholipids in high copy number which are exported into the host macrophage. The monoalkylglycerol of Leishmania lipophosphoglycan (LPG) and the Leishmania major GIPLs containing 1-alkyl-1,2-acylglycerol actively inhibit PKC in vitro [81]. The fatty acids were shown to be both competitive or noncompetitive inhibitors with respect to various PKC activators like dioeoylglycerol and phosphatidylserine [83]. Subsequently, Leishmania promastigotes deficient in lipophosphoglycan expression were found to invade macrophages but died soon after [84]. Loading LPG into the cells significantly prolonged their survival, and loading LPG into macrophages attenuated the ability of phorbol esters to induce the oxidative burst [84]. LPG and GIPLs were also associated with inhibition of oxidative burst, IL-1 production, chemotactic locomotion [85, 86], and downregulation of TNF receptors [82], all of which are PKC-dependent endpoints. These observations led to the hypothesis that Leishmania LPG has evolved to render macrophages anergic to activating signals. However, there are conflicting data on whether the carbohydrate or lipid domain of LPG is responsible for these activities. LPG is expressed primarily in the invasive promastigote stage and is downregulated in amastigotes, the chronic intracellular form of the parasite. Possibly, amastigote GIPLs exported into the host macrophage actually play a more important role in anergizing the cell to activation signals. A general preparation of amastigote GIPLs can antagonize interferon-웂-inducible iNOS expression, an activity shared by the highly purified iM4 GIPL of Leishmania [23]; the same molecule inhibits the translocation of PKC from cytosol to membrane (unpublished). Thus, when directly compared with GPIs of P. falciparum and T. brucei, the iM4 GIPL of Leishmania mexicana antagonizes PKC and NF-␬B/c-rel activation and downregulates the macrophage response to other stimuli, with opposing effects on other cell types [24], establishing specificity of action among different GPIs resulting from differences in fatty acid composition.


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NO has a major effect in killing Leishmania within macrophages [87], and interferon-웂 is the major physiological activator of iNOS in these cells. Thus, the activation of macrophages to a leishmanicidal state by interferon-웂 is a crucial mechanism in the immunological control of Leishmania infection. The findings that LPG and GIPLs of Leishmania are able to anergize macrophages to activating signals, that GIPLs in particular inhibit iNOS expression in response to interferon-웂, that GIPLs are exported into host macrophages, and that parasitelacking LPG are compromised in maintaining an intracellular infection in macrophages indicate that this class of glycoconjugate has evolved in Leishmania as a major mechanism of immune evasion [81, 82].

Regulation of Host Cells by Ceramide-Containing GPIs of T. cruzi There is evidence in T. cruzi infections of profound immunosuppression and derangement of T cell function, associated with a high proportion of apoptotic T lymphocytes [88–90], and parasite products are known to regulate T cell function [91, 92]. A recent study has shown that a ceramide-containing GPI derived from the lipophosphoglycan of T. cruzi profoundly suppresses activation of T cells through the TCR [93]. Ceramide-based signal transduction pathways have recently been shown to mediate a wide variety of cellular responses, especially cellular apoptosis [94–96]. These data therefore point to the regulation of T cells through apoptosis or anergy by T. cruzi ceramide-bearing GPIs. However, very large concentrations of glycoconjugates (in the range of 50–100 애M) were required; these exaggerated values raise the question of whether the effects were relevant. Ceramide alone at these concentrations is sufficient to mediate activity. It is not yet clear whether the GIPLs represent physiologic immunoregulators or simply a source of high concentrations of exogenous ceramide in these particular experiments.

Insulin-Mimetic Activities of Protozoal GPIs As noted above, there is circumstantial evidence for the involvement of GPIs in postreceptor insulin signaling. Malarial GPI is profoundly insulin-mimetic both in vitro and in vivo. When added at 1 애M to rat epididymal adipocytes in a standard assay of insulin activity, GPI was as effective as physiologically appropriate levels of insulin in inducing the uptake of [14C]glucose and conversion into triglycerides [70]. When administered to experimental animals, it induced a pronounced and prolonged hypoglycemia that was independent of TNF activity [70]. Other studies also show that T. brucei GPI is insulin-mimetic. The defined IPG of T. brucei MITat 1.5 inhibited isoproterenol-stimulated lipolysis at high dosage (100 애M) and suppressed hepatic gluconeogenesis by posttranslational modification of the two rate-limiting enzymes in gluconeogenesis, fructose-1,6biphosphatase and glucose-6-phosphatase [69]. The same authors later found evidence for the specific inhibition of isoproterenol-stimulated phosphorylation of intracellular substrates [97] at concentrations of 150 애M T. brucei GPI glycan.

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Exogenous GPIs also are insulin-mimetic [72, 98]: for example, 10 애M, the IPG fragment derived from the C-terminal GPI of human acetylcholinesteraseantagonized glucagon stimulation of glycogen phosphorylase. The clear inference from these studies, and the extensive literature implicating GPIs as a component of postreceptor insulin signaling [50–52, 60–62], is that parasite GPIs may induce hypoglycemia by substituting for endogenous GPIs of mammalian origin and contribute by an as yet undefined mechanism to postreceptor insulin signaling. Malarial and trypanosomal GPIs may well mediate the profound hypoglycemia often seen in both infections, a prognostic indicator of poor outcome.

CD1d-Restricted NK T Cells and Immunity to GPI-Anchored Proteins CD4⫹ NK1.1⫹ T cells are unusual lymphocytes [99] which produce IL-4 and IFN웂 very rapidly and at high levels in response to TCR ligation [100, 101]. The kinetics and magnitude of cytokine output suggest that these cells may be crucial regulators of downstream TH1/TH2 differentiation. NK T cells exhibit a skewed V움 /V웁 TCR repertoire [102, 103], with predominant usage of V움14 and V웁8 [104], consistent with selection by a limited range of ligands. Thymic NK T cells are positively selected by cortical thymocytes expressing the non-major histocompatability complex (MHC)-encoded but MHC-class I-like molecule CD1d [105], and peripheral NK T cells have been shown to exhibit autoreactivity to CD1d, leading to the idea that these cells may function in the absence of associated antigen recognition [106, 107]. However, the related human CD1b and CD1c molecules can elicit cytolytic and IFN-웂 responses by presenting mycobacterial glycolipid antigens to CD8⫹ or to double negative T cells [108, 109]. Furthermore, murine V움14⫹ NK T cells have been shown to recognize synthetic 움-galactosylceramide in the context of CD1d [110]. Of great significance is the observation that murine CD1d in transfected human T2 cells constitutively associates with phosphatidylinositol-containing compounds that appear to be endogenous GPIs [111]. These studies therefore suggest that CD4⫹ NK T cells may participate in CD1d-restricted recognition of glycolipid antigens. To date, however, the natural ligand and functional significance of NK T cells in immune responses in vivo remain unclear. In a recent study we have shown that GPIs of parasite and host origin are the ligands recognized in a CD1-restricted manner by NK T cells [112]. Priming of mice with malaria parasites leads to a substantial increase in the frequency of NK T cells. This cell population contains, but is not limited to, cells expressing V움14 and V웁8. These cells proliferate when exposed to GPIs of P. falciparum, T. brucei, L. mexicana, and a synthetic GPI of self-origin (rodent brain Thy-1), and they produce high levels of IL-4 in response to these GPIs. These responses are CD1-restricted as they can be elicited from CD1-transfected antigen-presenting cells (APCs) but not from sham transfectants, and from class II⫺ / ⫺ but not from CD1.1/CD1.2⫺ / ⫺ APCs, and are blocked by a monoclonal antibody to CD1. The anti-GPI CD1/NK pathway regulates, to a significant degree, the IgG response to GPI-anchored surface antigen such as the CS protein of malaria


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sporozoites and mfVSG of T. brucei. Allogeneic bone marrow or thymic chimeras, which cannot respond to normal protein antigens, are able to respond to GPIanchored proteins with the production of levels of IgG similar to syngeneic controls [112]. Malaria sporozoites are able to elicit anti-CS IgG in class II⫺ / ⫺ mice, lacking both class II and class II-restricted conventional CD4⫹ T cells (these mice are incapable of mounting IgG responses to antigens). Conjugation of GPI to the nominal protein antigen ovalbumin enabled anti-ovalbumin IgG responses in class II⫺ / ⫺ mice. Anti-CS antibody responses in vitro could be blocked substantially by anti-CD1, and CD1.1/CD1.2⫺ / ⫺ mice had reduced IgG responses to sporozoites [112]. GPI moieties may therefore act as ‘‘universal’’ T cell sites through presentation by the nonpolymorphic CD1 restriction element. As noted above, GPI-anchored surface proteins occur frequently among the medically important parasitic and fungal taxa such as Plasmodium, Trypanosoma, Leishmania, Toxoplasma, and Candida. As for all other protein antigens, T-cell-dependent antibody formation to these molecules is generally assumed to be MHC class II restricted. These data indicate the existence of a novel pathway for the regulation of immune responses to GPI-anchored proteins. Clearly, the CD1/NK T cell pathway constitutes a novel immunological phenomenon with great practical and theoretical significance for parasite immunology and the development of vaccines. The observations that CD1-restricted NK T cells can recognize mammalian GPIs [112] and that endogenous GPIs associate with CD1 [111] raise the possibility of autoreactive NK T cells with specificity for the GPI anchors of numerous mammalian proteins. NK T cells may represent a highly unusual, persistent autoreactive pool that plays a central role in the regulation of autoimmune diseases [113].

Conclusions A considerable body of literature implicates mammalian GPIs as mediating a novel pathway of signal transduction in multiple tissues in response to a wide diversity of agonists. Most of the evidence is only circumstantial, because studies either do not use purified material or cannot distinguish between the contribution to cellular endpoints of GPI signaling as opposed to protein ligands, receptors, other participants, and indeed the purely physical properties of GPI molecules. The demonstration that GPIs and GIPLs of parasite origin are alone able to activate or downregulate specific signaling pathways in host cells clearly establishes that these molecules do act as signal transducers or antagonists. Structurally diverse GPIs show specificity of action, particularly with respect to their fatty acid domains, particularly evident in comparisons of the action of GIPLs from Leishmania spp. with the action of GPIs of malaria and T. brucei. The production of NO in response to host signals is the primary effector mechanism in the immunological control of Leishmania, which is an obligate parasite of macrophages. Rendering the macrophage anergic to activation is a central strategy of immune evasion by this pathogen. In contrast, both malaria and T. brucei are potent macrophage activators, causing systemic inflammation. It appears that in each case structurally divergent GPIs/GIPLs play a central role in these host/

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parasite relationships. Through antagonizing PKC, alkylacylglycerols of Leishmania anergize macrophages to those signals involved in host defense such as interferon-웂, and they may have further effects in immune evasion through downregulation of antigen presentation. GPIs of malaria and of T. brucei are potent macrophage activators, and this may account for many features of pathogenesis in these infections. They are also insulin-mimetic and may be responsible for the profound hypoglycemia. Recent studies demonstrate the existence of a novel branch of the immune system, CD1-restricted NK T cells, with specificity for glycolipid antigens. This pathway appears to play a central role in the regulation of autoimmune diseases and may also contribute to control of parasitic infections. Consistent with the ‘‘danger model’’ of pathogen-initiated immune responses, CD1-restricted immunity may be intermediate between the innate ‘‘pattern recognition’’ and adaptive immune systems. GPIs may yet prove to constitute a major portion of the natural ligands recognized by the canonical V움14/V웁8 TCR repertoire of this unusual lymphocyte population. Although much remains to be elucidated concerning the role of GPIs in the host/parasite relationship, in seems clear that these glycolipid signaling molecules will prove to play a central role in the regulation of host physiology and in both the innate and acquired immune systems.

Acknowledgments This work received support from the UNDP/World Bank/WHO Special Program for Research and Training in Tropical Diseases, the National Health and Medical Research Council, the Clive and Vera Ramaciotti Foundation, and the William Buckland Foundation.

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9 Synthesis of the GPI Anchor Daniel Sevlever,* Rui Chen,† and M. Edward Medof† †Institute of Pathology, Case Western Reserve University, Cleveland, Ohio 44106 and the *Department of Research, Mayo Clinic Jacksonville, Jacksonville, Florida 32224

Introduction Hydrophobic transmembrane polypeptides serve as membrane-anchoring structures for most mammalian cell surface proteins. Posttranslationally added glycosylphosphatidylinositol (GPI) units alternatively attach some mammalian surface proteins to the plasma membrane. Among this latter group of glycoproteins are ectoenzymes, adhesion molecules, receptors, complement regulatory proteins, and histocompatibility antigens. In addition to attaching certain proteins in mammalian cells, GPIs also anchor surface proteins in protozoan parasites and yeast. In all species, GPI-anchoring moieties share a common linear core structure consisting of EthN-P-6-Man-움12-Man-움1-6-Man-움1-4-GlcN linked 움1-6 to an inositol phospholipid [1]. GPI anchoring is probably an ancient way to link proteins to the cell surface because, in contrast to the situation in mammals, it is the most common mechanism by which lower eukaryotes tether their proteins to the plasma membrane. However, as will be discussed later in this chapter, there may be reasons why certain mammalian proteins retain this anchoring mechanism. In the following sections, we review progress achieved in the elucidation of the mammalian GPI-anchor pathway. These advances were based on techniques developed in the analysis of the parasite counterparts and on the exploitation of GPI-anchored defective mutant cells that accumulate intermediates at different steps of the pathway. That some of the GPI intermediates can now be chemically synthesized has provided an invaluable tool for the ultimate characterization of the enzymes involved in GPI-anchor assembly. PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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In the past few years, most of the genes in the mammalian GPI-anchor pathway and some of the genes in the yeast pathway have been cloned. As a result of this and other work on the organization of GPI-anchored proteins in the membrane, further information is emerging not only on the assembly of GPI intermediates, but also on the topology of the synthetic pathway.

GPI Biosynthetic Pathway The entire pathway leading to the synthesis of the GPI anchor resides in the endoplasmic reticulum (ER). Most of the biosynthetic steps can be recreated in vitro using total microsomes or purified ER membranes [2–4]. The sketchy available information on the enzymes involved in GPI-anchor assembly is derived primarily from these in vitro reactions. A proposed GPI biosynthetic pathway in mammalian cells is diagrammed in Fig. 9-1. This pathway is based on earlier studies of GPI biosynthesis in the trypanosome Trypanosoma brucei [5, 6] and on analyses of free GPIs, i.e., GPIs unlinked to proteins which accumulate in

Figure 9-1 Steps and structures of mammalian intermediates of the biosynthesis of the mammalian GPI-anchor precursor. Transfer of GlcNAc to PI initiates the assembly of the GPI anchor. After deacetylation of GlcNAc and acylation of inositol, sequential addition of Man and EthN-P to GlcN-(acyl)PI follows. The donors of palmitic acid for inositol acylation and EthN-P to the first and second mannosyl residues have not been conclusively established.

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several mammalian mutant cell lines that are deficient in cell-surface GPIanchored protein expression [4, 7–10]. The pathway starts with the transfer of N-acetylglucosamine (GlcNAc) from UDP-GlcNAc to phosphatidylinositol (PI) by the UDP-GlcNAc : PI transferase [5]. This enzyme may recognize a select pool of PI, because available data indicate that the lipid groups of the glycerol backbone of GlcNAc-PI contain a higher percentage of alkyl chains than in most common PI species [4]. The second step consists of the deacetylation of GlcNAc-PI into glucosaminyl-phosphatidylinositol (GlcN-PI). Deacetylation is followed by the addition of palmitic acid to the inositol ring [11, 12]. Inositol acylation is an obligatory step in mammalian cells and yeast (see below), and renders GlcN-(acyl)PI and further downstream mammalian intermediates resistant to phosphatidylinositol-specific phospholipase C (PI-PLC) hydrolysis, the most widely used tool to assess GPI anchoring (see Chapter 11 by Low). Two different mechanisms, one dependent on acyl-CoA and the other acyl-CoA independent, have been reported for this fatty acid addition to GlcNPI, the choice of which may be a function of the nature of the substrate offered. When endogenous GlcN-PI is the substrate in in vitro experiments, the acylation is acyl-CoA independent [13]. Conversely, the reaction becomes acyl-CoA dependent when chemically synthesized GlcN-PI is used [14]. The latter mechanism for GlcN-PI acylation in mammals is in accordance with that reported in yeast [15]. To this point, the mammalian and yeast pathways are identical. They differ, however, from the biosynthesis of the GPI anchor in the bloodstream T. brucei parasite where inositol acylation is not essential for the pathway to proceed [16, 17]. Inositol acylation appears to have different roles in mammalian cells and parasites. In parasites, GPI-anchor intermediates and the fully assembled precursor can be found in two versions, nonacylated and acylated, and only the nonacylated precursor appears to be transferred to proteins [18]. Because both precursors are in dynamic equilibrium, the acylated GPI may serve as a reservoir of anchor precursors in T. brucei [19]. In mammals, inositol acylation seems to be necessary for efficient mannosylation and transfer of the fully assembled GPI donor to proteins (see below). Following inositol acylation in mammalian cells and yeast, GlcN-(acyl)PI serves as a substrate for the addition of the first of the three mannosyl residues, all of which are donated by dolicholphosphomannose (DPM). Glycosylphosphatidylinositol mannosyl-transferase-I (Gpimt-1p) can utilize either DPM or mannosyl-phosphopolyprenol (the donor of Man residues in prokaryotes) in in vitro systems, but the rate of Man-GlcN-(acyl)PI formation is twofold higher with DPM [20]. Gpimt-1p recognizes the acyl group on the inositol ring as evidenced by the fact that GlcN-PI cannot serve as a substrate for the enzyme [14, 21]. In mammals, the main pathway continues with the addition of ethanolaminephosphate (EthN-P) to the initially transferred Man residue (Man1), although mannosylated GPIs species without EthN-P attached to the first mannosyl residue also have been identified [22]. Modification of Man1 by EthN-P is a hallmark of mammalian GPI-anchor structures. Recently, this modification also has been uncovered in GPI-anchor intermediates from yeast [23], but it is not clear whether it is also present in protein-associated GPI anchors in yeast as is the case

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in mammals [1]. Phosphatidylethanolamine is the donor of at least one of the EthN-P groups incorporated into mammalian GPIs [24]. The second and third mannoses (Man2 and Man3, respectively) of the GPI core glycan also are transferred from DPM [25, 26]. Addition of the third mannosyl residue can be inhibited by mannosamine [27–29]. This amino sugar is one of the few inhibitors of mammalian GPI-anchor synthesis that is active on addition to intact cells [29]. Recently, a natural compound isolated from yeast has also been found to inhibit the 움1-2 linkage between Man2 and Man3 in mammals and yeast but not in protozoan parasites [30]. Transfer of the third mannosyl residue is followed by its modification by the EthN-P that provides the link between the fully assembled GPI anchor and the protein. The addition of this terminal EthN-P can be blocked with phenylmethylsulfonyl fluoride in T. brucei parasites but not in mammalian cells [31, 32]. The complete GPI, termed H7 [22], that results from the addition of the terminal EthN-P can be further modified by the attachment of a third EthN-P group to Man2 to produce H8. Based on structural similarities between the last two intermediates H7 [EthN-P-Man-Man(EthN-P)-Man-GlcN-(acyl)PI] and H8 [EthN-P-Man-(EthN-P)-Man-(EthN-P)Man-GlcN-(acyl)PI] to the GPIs found attached to proteins, H7 and H8 are the most likely GPI-anchor donor candidates [22, 33]. This proposition is further supported by the high turnover rate of H7 [34]. However, in order to reconcile the structures of H7 and H8 with the GPIs attached to proteins, one has to postulate lipid remodeling reactions before, during, or after transfer to the protein. Among these, deacylation of the inositol ring is particularly important because the vast majority of GPI-anchored proteins, with the exception of those on human erythrocytes, do not contain this modification and hence are susceptible to PI-PLC hydrolysis [35]. The existence of a deacylase was initially suggested by the demonstration that the PI-PLC resistant phenotype behaves as a recessive trait in stable hybrids prepared from cell lines expressing PI-PLC resistant and PI-PLC susceptible GPI-anchored proteins [36]. Recent studies have shown that the transfer of acylated GPI-anchor donors is followed by the acquisition of PI-PLC susceptibility of the GPI in newly synthesized protein [37]. This conversion of the GPI anchor in GPI-anchored proteins from PI-PLC resistant to PI-PLC susceptible takes place in the ER [37]. Based on these data, the likely sequence of events is first the transfer of an acylated GPI anchor to a protein followed by the rapid deacylation of the transferred GPI. A second type of lipid remodeling has been proposed [34, 38]. Diacyglycerol, present to varying degrees in the GPI-anchor intermediates, replaces alkylacylglycerol as found in the GPI anchor of mammalian proteins [erythrocyte acetylcholinesterase, decay-accelerating factor (DAF), and placental alkaline phosphatase] analyzed to date [35, 39, 40]. A notable exception is CD52 from human lymphocytes, which contains diacylglycerol in its GPI anchor [41]. Although supporting data for the above proposed remodeling is still lacking, some progress in understanding the origin of the alkyl chain present in the GPI anchor of certain proteins [34, 38, 42, 43] has been made. Analysis of the GPI anchors of proteins in cells deficient in the peroxisomal enzymes involved in the synthesis of ether-linked glycerolipids has shown that levels of alkylacylglycerol are unchanged in the glycerol backbones of GPI anchors, suggesting that these lipids probably do not derive from peroxisomes [38].


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The pathway described above for mammals is broadly similar to that in yeast and in the T. brucei parasite. However, as indicated above, in the mammalian and yeast pathways, inositol-acylated species predominate. In contast, in T. brucei, nonacylated species are the abundant intermediates, and lipid remodeling occurs at the level of the fatty acids in the fully assembled anchor prior to protein transfer [17]. In the yeast pathway, diacylglycerol is replaced by ceramide after anchor transfer (Fig. 9-2) [44], which more closely parallels the mammalian pathway [37]. The first two steps of the GPI biosynthetic pathway take place on the cytoplasmic face of the ER [45]. Support for this proposition has been provided by topological studies of GPIs and the gene products responsible for the addition of GlcNAc to PI and the deacetylation of GlcNAc-PI [45–47], see next section. In T. brucei parasites, the remainder of the steps of GPI-anchor synthesis has also been reported to take place on the cytoplasmic face of the ER [48]. Although due to the lack of suitable probes, no comparable data are available on the mammalian counterparts, the ER orientation of the downstream PIG-B gene product (see below) suggests an intraluminal orientation. In mammalian cells, all GPI-anchor species are not always completely consumed in anchoring proteins, and at least some unused free GPIs can exit the ER and reach the plasma membrane. Evidence for this has been obtained using three different approaches: (a) labeling of GPIs present in the outer leaflet of the plasma membrane with an impermeant biotinylated probe [49]; (b) recovery of GPIs from budding viral particles [50]; and (c) coisolation of GPIs with membranes enriched in plasma membrane markers [34]. Interestingly, although GPI anchors are thought to serve as sorting signals for the apical localization of GPI-anchored proteins in polarized epithelia (see below), free GPIs in one study [50] were found to be present to the same extent on both apical and basolateral surfaces. Because tight junctions on the outer plasma membrane leaflet of polarized epithelial cells would preclude diffusion to the basolateral surface on the outer leaflet, this result would require free GPIs to be present on the inner leaflet. The topology of free GPIs at the plasma membrane, therefore, is still controversial.

Mammalian and Yeast GPI-Anchor Defective Mutants The derivation of Thy-1 negative murine lymphoma mutant cell lines [51], which occurred long before the GPI moiety was postulated to be an alternative anchor for cell surface proteins, was instrumental in the identification and characterization of most of the GPI-anchor intermediates. These lymphoma mutants fall into six complementation classes [52]. Five of these mutants make normal amounts of Thy-1 polypeptide but fail to express the mature protein at their cell surface. Classes A, C, and H are defective in the first step of GPI synthesis, the transfer of GlcNAc to PI (see Fig. 9-1). The genes that encode the proteins affected in classes A, C, and H cells, PIG-A (for phosphatidylinositol glycan class A) [53], PIG-C [54], and PIG-H [55], have been isolated by expression cloning. Mutations in the gene termed PIG-A [56] have been shown to be responsible for the GlcNAc-PI synthase defect that is present in hematopoietic stem cells from

Figure 9-2 Postulated lipid remodeling pathways of GPI anchors. GPI anchors in mammalian cells exist in two pools: (1) a large pool with a diacylglycerol substituent and (2) a minor one with alkylacyglycerol. These two pools may exist in equilibrium as indicated by the broken arrow. Preliminary data (Chen and Medof, unpublished) indicate that after transferring to proteins the diacylglycerol is replaced by alkylacylgyerol in the already attached GPI anchors (these putative pathways are indicated by thicker arrows). Alternatively, the minor pool of alkyacylglycerol GPI anchors could be preferentially transferred to proteins. The other type of remodeling involves deacylation of the inositol and takes place in the GPI anchor soon after its attachment to protein [37]; this reaction appears to occur before the above postulated lipid replacements (Chen and Medof, unpublished). The situation in yeast cells is different, where GPI-anchored proteins can contain two different type of lipids: diacyglycerol (the only known protein containing this lipid is Gas1p, the most abundant GPI-anchored protein in yeast) or ceramide. Yeast GPI-anchor precursors appear to contain only diacylglyerol when they are transferred to


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patients with the acquired hemolytic anemia, paroxysmal nocturnal hemoglobinuria (PNH) [57, 58] (see Chapter 2 by Luzzatto and Nafa). Three Saccharomyces cerevisiae mutants, termed gpi-1, gpi-2, and gpi-3, are also defective in the first step of GPI assembly [59, 60]. The corresponding yeast proteins Gpi2p and Gpi3p, are homologous to the mammalian PIG-C and PIGA gene products Pig-Cp and Pig-Ap, but Gpi1p shows no homology with PigHp. Recently, the mammalian homologue of GPI1 [61, 62] was cloned and its product Gpi1p shown to form a complex with Pig-Ap, Pig-Cp, and Pig-Hp [62]. The isolated complex, comprised of these four gene products, appears to be sufficient to carry out the first step of GPI anchor synthesis [62]. Topology studies of Pig-Ap have revealed (1) a large cytoplasmic amino terminal domain where the catalytic activity is presumed to reside (based on its similarities to a bacterial GlcNAc transferase), and (2) a small carboxy-terminal domain [46]. Pig-Hp has been proposed to have a hairpin structure with both its amino and carboxy termini facing the lumen of the ER [46]. Mutants deficient in the second step of the pathway are not represented within the set of Thy-1 negative lymphoma mutants. These mutants, blocked in the deacetylation of GlcNAc-PI (see Fig. 9-1), were first isolated in K562 cells [63] and termed class J, and more recently in CHO cells [47, 64] and termed class L. It is not known whether the defect in these two mutants is the same, as complementation studies between the two have not been done. The gene named PIG-L that rescues the CHO mutant has also been shown to be oriented in the ER toward the cytoplasm [47]. The membrane orientation of the gene products involved in these first two steps of the pathway is consistent with topology studies of the GPI intermediates GlcNAc-PI and GlcN-PI using PI-PLC which concluded that both GPIs are facing the cytoplasm [45]. No mutant affecting GPIMT-1, the gene which is responsible for carrying out the fourth step of the pathway, is available as yet. However, lack of synthesis or availability of the DPM substrate can block the pathway causing the accumulation of GlcN-(acyl)PI, as is the case in lymphoma complementation class E [51, 12] and the CHO mutant Lec 35 [65], respectively. GlcN-(acyl)PI accumulation has also been documented in the HeLa S3 subline, despite the expression of normal levels of DPM and the ability of the cells to use this substrate to fully mannosylate downstream GPIs intermediates and N-glycans [66]. Downstream in the pathway, the next mutant available is Thy-1⫺ lymphoma class B, which is defective in the addition of the third mannosyl residue and as a consequence accumulates Man-(EthN)-Man-GlcN-(acyl)PI [67]. PIG-B, the

the nascent polypeptide. The GPI anchor then can follow two alternative routes. (1) The diacylglycerol remains but the fatty acid at the sn-2 position is replaced by a longer C26:0 [124] or (2) the diacylgylcerol is replaced by ceramide as is probably the case for the majority of yeast GPI-anchored proteins [44]. Further exchange of the initially incorporated ceramides with more polar ceramides has been proposed [125]. Yeast GPI anchors do appear to undergo deacylation of inositol as in mammals: despite PI-PLC resistance of GPI precursors, yeast GPI-anchored proteins are susceptible to the enzyme [44]. These steps are indicated by broken arrows. Finally, in T. brucei parasites the initially more hydrophobic fatty acids at the sn-1 and sn-2 positions of the GPI-anchor donor are replaced sequentially by myristic acid before transfer to variant surface glycoproteins (VSGs) [17]. Lyso-acyl intermediates bearing one myristic acid have been detected [17]. Myristic acid exchange in the GPI anchor after transferring to VSGs also has been recently proposed [126].

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gene that restores normal GPI anchoring in lymphoma mutant B cells, has been cloned and encodes an ER protein with a relatively short N-terminal domain accessible from the cytoplasm and a long carboxy-terminal domain inside the ER lumen [68]. PIG-B constructs lacking part of the N-terminal domain are still able to restore GPI anchoring, indicating that this portion of the molecule is not needed for enzymatic activity and further suggesting that the active site resides in the luminal domain [68]. The location of the Pig-Bp active site in the ER lumen provides a strong indication that the transfer of the third mannosyl residue from DPM to the GPI occurs on this side of the ER membrane. The recent cloning of a yeast functional homologue of PIG-B, GPI10, underscores the highly conserved GPI biosynthetic machinery throughout evolution [69]. Lymphoma mutant class F accumulates Man-Man-(EthN-P)-Man-GlcN(acyl)PI (termed H6, see Fig. 9-1) as a consequence of being defective in the addition of the terminal EthN-P to the third mannosyl residue [8]. This EthNP group attached to Man3 provides the bridge between the GPI anchor and the polypeptide. PIG-F cDNA complements the defect of class F mutant cells and encodes a putatively very hydrophobic protein [70]. No information on the topology of the PIG-F gene product is available. Cells deficient further downstream in the machinery that transfers anchor donors to proteins are expected to accumulate complete GPIs as well as incompletely processed proteins retaining their C-terminal signal peptides. The phenotype is exhibited by a mammalian mutant cell line [71] and two yeast mutants [72, 73]. The mammalian mutant cell line, a human K562 derived line termed class K, accumulates H7 and H8, the two terminal ethanolamine-phosphatesubstituted putative GPI donors, but fails to display GPI-anchored proteins at its cell surface [71]. The two yeast temperature-sensitive mutants that exhibit the same phenotype are termed gpi-8 [74] and gaa-1 [73]. Mutant gpi-8 accumulates one of the complete yeast GPI lipid donors CP2 (see below) which normally remains undetectable [74]. Cloning of the defective gene, yGPI8, in gpi-8 cells has shown that it encodes a 411 amino acid-long ER protein. yGpi8p is a type I integral membrane protein with a large lumenally oriented N-terminal domain and short C-terminal cytoplasmic sequence. When transfected with yGPI8 cDNA, the growth defect and Calcofluor white hypersensitivity of gpi-8 cells are abolished [74]. Similar to the gpi-8 mutant, at restrictive temperature the other yeast mutant with the same phenotype, gaa-1, accumulates both of the complete yeast GPI-anchor donors, CP1 and CP2, but does not attach either donor to proteins [73]. Cloning of the defective gene, yGAA1, in the gaa-1 line has shown that it encodes a 614 residue ER protein. Reconstitution of the gaa-1 mutant with yGAA1 cDNA improves its ability to attach anchors to a recombinant Gas1p protein bearing mutated anchor attachment sites (termed 웆, see next section). yGaa1p has a large, approximately 306 amino acids long, lumenal domain with two N-linked glycans and five membrane spanning domains. At the extreme C terminus of the protein there is an ER retrieval signal (KEKQS) [73]. The murine homologue of yGAA1, mGAA1, has been cloned from a 6-day embryoid body cDNA library by the signal-sequence trap method [75]. The human homologue, hGAA1, has been cloned by probing a human fetal heart cDNA library with the murine sequence. hGAA1 cDNA is approximately 2 kb


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in length and like yGAA1 encodes a 621 amino acid long protein. The amino acid sequence of hGaa1p is 25% identical and 57% homologous to that of yGaa1p. Kyte-Doolittle hydrophobicity plots of both proteins show marked similarity [75]. The human homologue of yGPI8 has been cloned by 5⬘ extension of a partial Genbank sequence showing homology to the yeast cDNA [76]. hGPI8 encodes a 395 amino acid long glycoprotein that shows 43% overall identity with yGpi8p. hGPI8 gives rise to mRNAs of 1.6 and 1.9 kb, both of which encode the same 395 amino acid long protein (45 kDa). The levels of both mRNAs correlate in cells with their ability to couple GPIs to proteins. The gene spans 앑2.5 kb of DNA on chromosome 1 [76]. hGpi8p has a similar hydrophilicity profile to yGpi8p. RT-PCR analysis of endogenous hGPI8 mRNA in class K cells has shown the presence of two mutations: deletion of an A105T106 which results in a stop codon (TAG) at position 131, and deletion of one of six tandem T residues 앑300 bp downstream of this position. Transfection of class K cells with hGPI8 cDNA abolishes accumulation of GPI precursors and restores C-terminal processing of GPI-anchored proteins, resulting in their normal expression. These data, in conjunction with in vitro studies of microsomes from K cells employing a translation processing system, have indicated that this gene encodes the transamidase enzyme (see next section). No natural mammalian cell mutant defective in the GAA1 gene has been identified. Genomic cloning of the mGAA1 gene has shown that it contains 12 exons and 11 introns [75, 77]. Mapping of a YAC library with hGAA1 cDNA has shown that the gene spans 40 kb at q12-13 of chromosome 2 in humans [78]. With the availability of murine genomic [77] sequence as well as mGAA1 cDNA sequence [75], both antisense mutants in 3T3 fibroblasts and recombinant knockouts in F9 cells [77, 79] have been prepared. Biosynthetic labeling of the mutant cells has shown that they accumulate H7 and H8 as do mutant K cells.

GPI-Anchor Addition to Proteins Proteins that are destined to be GPI anchored are synthesized with two hydrophobic signal peptides, both of which are removed during posttranslational processing: (1) an amino terminal peptide to direct the protein to the ER and (2) a carboxy-terminal GPI-signal sequence. The latter peptide consists of between 17 to 31 residues that are recognized by the GPI transamidase complex. This signal sequence contains three elements that are required to allow GPI attachment: (1) a hydrophobic domain, (2) a cleavage/attachment site termed 웆 that is located 10–12 residues N terminal to the hydrophobic domain, and (3) a spacer, the previously described region between the cleavage/attachment site and the hydrophobic domain (reviewed in [80]). Comparisons of GPI-signal sequences among GPI-anchored proteins have not revealed a consensus sequence, and rather only a predominance of hydrophobic amino acids is evident; substitutions in the carboxy terminal GPI-signal sequence are tolerated as long as the general hydrophobicity is maintained. Furthermore, GPI-signal sequences of any given GPI-anchored protein are functional when exchanged with that of another. The cleavage/attachment site consists of the 웆 residue, to which the ethanolamine amino group of the GPI anchor is attached, and the two adjacent down-

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stream amino acids (웆 ⫹1 and 웆 ⫹2) [81]. The amino acids which function at the 웆 site are restricted to serine, glycine, alanine, aspartic acid, cysteine, and asparagine (see Fig. 9-3) [81, 82]. A common feature in these amino acids is the presence of a small side chain. There is less stringency for the 웆 ⫹1 residue, but the same general rule applies for the 웆 ⫹2 amino acid. Studies of the yeast protein Gas1p similarly have found that only small amino acids at or within the cleavage/attachment site are compatible with efficient GPI addition [83] (Fig. 9-3). No selectivity appears to reside in amino acids N terminal of the 웆 site: 웆 ⫺1, 웆 ⫺2, etc. Finally, substitutions, deletions, and additions within the spacer region of mammalian and yeast proteins indicate that both the distance and the sequence between the hydrophobic domain and the 웆 site are important [83]. Failure of GPI-anchor addition to proteins can occur due to the inability of the cells to synthesize GPI anchors or to the presence of uncleavable sequences at or around the 웆 site. Initially, the fate of Thy-1 protein was analyzed in Thy1⫺ lymphoma mutants with defects at different sites in the GPI biosynthetic pathway [84, 85]. Later, studies were done in which the 웆 site was mutated [82, 86, 87]. Taken together, these studies indicated that irrespective of the way in which GPI anchoring is blocked, the consequences for the protein seem to be the same. It stalls in an early compartment of the secretory pathway, probably the ER or an ER–Golgi intermediate compartment, where it forms disulfidelinked aggregates and then is transported to the cytoplasm (see below) and degraded [87]. The GPI-signal sequence thus may fulfill a dual role, as a recognition signal for the transferase for GPI addition or as an ER degradation signal. Studies of the mechanism(s) involved in the degradation of a protein containing an uncleavable GPI signal have shown that inhibitors of lysosomal activity have no effect on the processing of the polypeptide [87]. However, specific inhibitors

Figure 9-3 Amino acid requirements of the GPI-transamidase complex. Using the signal sequences of a mammalian and a yeast GPI-anchored protein, DAF and Gas1p, respectively, the amino acids tolerated by the GPI transamidase complex at the 웆 and 웆 ⫹2 sites are shown.


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of cytosolic proteasomes greatly reduce degradation, suggesting the possible involvement of this pathway when GPI addition is impaired [88]. It has been postulated that the enzyme responsible for the removal of the GPI-signal sequence also transfers the GPI moiety to the protein in a transamidation mechanism. The putative transamidase has been studied in an in vitro translation system using microsomal membrane preparations as a source of enzyme and the GPI anchor, and an engineered form of placental alkaline phosphatase cDNA (termed miniPLAP) as the cDNA encoding the protein-substrate (see Fig. 9-4). The engineered PLAP sequence, which is devoid of the glycosyla-

Figure 9-4 Various outcomes of COOH-terminal GPI-signal peptide processing as assessed using the miniPLAP in vitro translation system. Following translation, the primary translation product translocates into the ER lumen and undergoes N-terminal processing in which the NH2 signal (checkered bars) is cleaved. In the miniPLAP system, this can be monitored by the conversion of 28-kDa preprominiPLAP to 27-kDa prominiPLAP (inset, lower left). Under normal conditions and for most of this product, the COOH-terminal signal peptide (diagonally hatched bars) is subsequently cleaved and a GPI-anchor donor substituted (central pathway). In the miniPLAP system, this is reflected by the appearance of 24.7-kDa mature miniPLAP (inset, lower left). In normal cells, for a small amount of the NH2terminal processed product, or in GPI-anchor-defective cells in the absence of sufficient GPI-anchor donors, the COOH-terminal signal peptide is cleaved and water is added in place of the GPI donor (upper pathway). In the miniPLAP system, this is reflected by the presence of 23-kDa ‘‘free miniPLAP’’ (inset, lower left). A product of identical size is generated if the alternative nucleophiles HDZ or HDX are added. If the nascent polypeptide contains a defective site [81], it undergoes N-terminal processing but does not undergo subsequent C-terminal signal peptide cleavage (lower pathway). In the miniPLAP system, this is reflected by the accumulation of 27-kDa prominiPLAP (inset, lower left). The incompletely processed propeptide is retained in the ER and degraded (not shown, see text).

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tion sites and the catalytic domain, allows monitoring of small changes in molecular weight that take place during processing of the N- and C-terminal domains [89] (see Fig. 9-4). Studies in this translation system have shown [90] that ATP and GTP stimulate N- and C-terminal processing of the nascent protein. This energy requirement initially appeared inconsistent with the proposition that a transamidase is responsible for the GPI addition to proteins. However, subsequent studies showed that the uncleaved nascent protein interacts with the ER chaperone BiP and that ATP is required for this interaction, not for the formation of the amide bond [91]. The association of BiP with an unprocessed putative GPI-anchored protein containing a mutated 웆 site has also been demonstrated in cells [92]. Thus, it appears that the energy-dependent reactions precede the transamidase reaction. Evidence that the condensation between the amino group of EthN of the GPI anchor and the carboxyl group of the amino acid at the 웆 site occurs through a transamidation has been obtained using the small nucleophiles, hydrazine (HDZ) and hydroxylamine (HDX) [93] (see Fig. 9-4). HDZ and HDX have been shown to act as nucleophilic acceptors in transpeptidase and transamidase reactions [94]. This implies that HDZ and HDX are able to generate an activated carbonyl group, a hallmark of a transamidase, which is then attacked by the ethanolamine. The microsomal system has also been useful in determining the nature of the defect in mutant K cells. When microsomes from mutant K cells were used instead of those from wild-type cells, N-terminal processing of miniPLAP was documented. However, unlike other mutants affected at upstream GPI assembly steps, when mutant K microsomes were assayed, they showed no detectable Cterminal processed minPLAP irrespective of reaction conditions [95]. Further experiments demonstrated that, in distinction to all other GPI anchoring mutants described, mutant K microsomes failed to support C-terminal signal peptide replacement by HDZ or HDX [95]. In contrast, following transfection with hGPI8 cDNA, microsomes from K cells were rendered competent in using HDZ and HDX as nucleophilic donors (see previous chapter) in place of the GPI anchor [76].

Biological Implications of GPI Anchoring Probably the only undisputed function of GPI anchors is to provide the attached polypeptides with a stable association with the outer plasma membrane leaflet. While many other roles for the anchor have been proposed, none have been conclusively supported by experimental data. Higher lateral mobility in the plane of the plasma membrane as a GPI-anchor property has been suggested because GPI-anchored proteins cannot directly interact with the cytoskeleton because they do not cross to the inner leaflet of the plasma membrane and certain GPIanchored proteins appear to exhibit higher lateral mobility than membranespanning proteins [96]. However, lateral mobility measurements obtained using chimeric proteins of membrane-spanning and GPI-anchored proteins have concluded that the major determinant of the high mobility of the latter proteins resides in their extracellular domains, and the contribution of the GPI anchor


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to the higher mobility displayed by many GPI-anchored proteins [96] is less important. Another role for GPI anchoring could be to provide for the release of the attached proteins under certain circumstances. Soluble forms of GPI-anchored proteins have been detected, although the mechanisms that generate these forms remain unknown. The GPI anchor is susceptible not only to hydrolytic enzymes like bacterial and parasite PI-PLC, but also to a more specific enzyme, mammalian serum GPI-specific phospholipase D (GPI-PLD). However, in contrast to bacterial PI-PLC, GPI-PLD is unable to cleave GPI-anchored proteins in intact cells [97–99]. Interestingly, at least in some cell lines (e.g., HeLa and myeloid cells), the presence of an endogenous GPI-PLD that is able to release GPI-anchored proteins into the media has been reported [100, 101] and possibly some of the GPI-anchored proteins found in extracellular fluids may be generated in this way. GPI anchors might target proteins to particular domains either in the secretory pathway on their way to the plasma membrane, while at the plasma membrane, or after recycling from it. GPI-anchored proteins have been shown to associate early in the secretory pathway (probably in the Golgi apparatus) with specialized membrane domains [102] that contain cholesterol, sphingolipids, and other cytoplasmic lipid-modified proteins like nonreceptor type tyrosine kinases and heterotrimeric G proteins [103] (see Fig. 9-5). The complexes remain intact upon their arrival at the plasma membrane. The membrane domains can be isolated by virtue of their resistance to Triton X-100 solubilization at 4⬚C and by their buoyant density in sucrose gradients [102]. Triton insoluble membranes (TIMs) is one among the many acronyms that have been used to describe these structures. Some of the biological properties of GPI-anchored proteins (see below) can be explained in the context of their presence in TIMs. A further proposed function of GPI anchors is that in polarized epithelia, they direct their anchored polypeptides to the apical surface and as a result concentrate GPI-anchored proteins preferentially at this site [104, 105]. Replacement of the transmembrane domain of proteins that are normally expressed basolaterally with a GPI anchor will reroute them to the apical surface [105] and, based on these results, the GPI anchor has been postulated to be an apical sorting signal [106]. Apical sorting may be the default pathway, and therefore it is possible that the lack of a transmembrane domain containing basolateral signals rather than the presence of the anchor in GPI-anchored proteins causes them to go to the apical surface. Because sorting decisions are made in the trans-Golgi, where apparently the association of GPI-anchored proteins with TIMs occurs, it has been proposed that TIMs might serve as rafts to ferry molecules destined to go to the apical surface [103]. However, in MDCK cells, TIMs have been shown to exist on both the apical and basolateral surfaces [107, 108]. Interestingly, in spite of the ability of GPI-anchor intermediates to associate with TIMs [109], free GPIs are apparently distributed on both the apical and basolateral surfaces of polarized cells [50]. This finding also appears to contradict the proposition that the GPI anchor is alone responsible for the protein targeting to TIMs that results in the apical localization of GPI-anchored proteins in polarized epithelia.

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Figure 9-5 Schematic representation of Triton insoluble membranes (TIMs) of the plasma membrane. Although TIMs can occur in intracellular compartments [127], only the known components of plasma membrane TIMs are depicted in the figure. The thicker lines represent plasma membrane ‘‘patches’’ or TIMs where GPI-anchored proteins, free GPIs, sphingomyelin, glycosphingolipids, cholesterol, non-receptor-type tyrosine kinases, and a few transmembrane proteins are present. Not all of these proteins are constitutively present in TIMs but are recruited into these domains upon stimulation (antibody or ligand binding). TIMs are believed to encompass both leaflets of the plasma membrane; only non-receptor-type tyrosine kinases, heterotrimeric G proteins, cholesterol, and probably free GPIs are known to exist in the inner leaflet. A common theme of the proteins present in TIMs is that they are modified by lipids: GPI, fatty acids, and/or prenyl groups (reviewed in [128]). However, it has been reported that in two lipid-modified proteins, hemagglutinin (HA) and caveolin, the lipid is not responsible for their presence in TIMs [113, 129]. Some transmembrane proteins without lipid modifications also have been reported to be present in TIMs.

Recent evidence has implicated GPI anchors in the relatively longer time that GPI-anchored proteins (앑three times higher than the transferrin receptors and C6-NBD-sphingomyelin) spend in early endosomes before recycling back to the cell surface [111]. Lowering membrane cholesterol levels has been shown to increase the rate by which GPI-anchored proteins return to the plasma membrane. Because TIMs appear to be disrupted when cholesterol is lowered (using either inhibitors of its synthesis and/or cholesterol binding agents) [112, 113], it has been proposed that the endosomal retention of GPI-anchored proteins is due to their presence in TIMs [111]. As an extension of this hypothesis, it has further been proposed that this retention of GPI-anchored proteins in early endosomes could explain why only the GPI-anchored prion protein and not its transmembrane chimera can be converted to the infectious scrapie form by a process which is cholesterol dependent [114]. Another proposed property of GPI-anchored proteins is that they can elicit transduction signals upon antibody cross-linking despite their lack of intracellular


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domains (reviewed in [115, 116]). In several instances, signaling function has been shown to reside on the anchor, because transmembrane versions of these proteins lose this capability [117]. Antibody cross-linking of GPI-anchored proteins induces rapid tyrosine phosphorylation of certain proteins, suggesting the involvement of tyrosine kinases [117]. GPI-anchored proteins can be coimmunoprecipitated with non-receptor-type protein kinases and both components are present together in TIMs (reviewed in [115]). This suggests, albeit indirectly, that although they reside on different leaflets of the plasma membrane, GPIanchored proteins and non-receptor-type protein kinases are in close proximity presumably within TIMs. Finally, purified GPI-anchored proteins have been shown to possess the ability to spontaneously reintegrate into cell membranes (reviewed in [118, 119]). Intercellular transfer of GPI-anchored proteins has been demonstrated in HeLa cells [120] and in transgenic animals [121]. In at least one instance this phenomenon has been shown to have physiological consequences: the avoidance by Schistosoma mansoni parasites of complement-mediated lysis by acquiring complement protecting GPI-anchored proteins (i.e., DAF) from their host [122]. The spontaneous transfer of GPI-anchored proteins can be exploited to reengineer cell surfaces with GPI-anchored proteins, a technology termed ‘‘painting’’ [118], as an alternative to gene transfer. Potential applications of this technology include anticancer vaccines, protection against graft rejection, downregulation of autoimmune responses, and in vivo targeting among others [118].

Conclusions In the past decade, a great deal has been learned of how the mammalian GPI anchor is synthesized and attached to proteins. Most of the genes involved in the pathway have been cloned and some insight on the characteristics of the activities encoded by these genes is starting to emerge [123]. Among other things, this knowledge may have potential therapeutic implications, as differences between the mammalian, yeast, and parasite pathways are becoming apparent. Furthermore, seminal studies on the ability of the GPI anchor to target polypeptides to discrete plasma membrane areas not only opened a new field of research, but also provided a rational for the puzzling ability of GPI-anchored proteins to participate in signaling cascades. Lagging behind, however, has been progress in understanding the role of the GPI-anchor intermediates in the plasma membrane and in revealing the mechanisms by which these glycolipids travel from their site of synthesis to the cell surface. Generating probes that recognize GPIanchor intermediates will be helpful in advancing our current knowledge in this area of research.

Acknowledgments This work was supported by NIH grants AI23598 and HL55773 (EM) and DK55002 (DS). The authors thank T. Rosenberry for critical reading and Sara Cechner for manuscript preparation.

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10 Functional and Structural Organization of GPIAnchored Proteins in Cellular Membranes Satyajit Mayor National Centre for Biological Sciences, Hebbal, Bangalore 560 065, India

Introduction Glycosylphosphatidylinositols-anchored proteins (GPIAPs) are a large class of proteins of extreme diversity in structure and function, and their sole common feature is that they all share virtually identical glycolipid membrane anchors. GPI addition is a posttranslational modification that serves as a membrane anchor and involves the replacement of the carboxyl-terminal peptide sequence of the protein by a glycosyl-inositol phospholipid moiety [1–5]. GPI anchors have an evolutionarily conserved core glycan that bridges an ethanolamine residue in amide linkage with the protein and an inositol phospholipid [4–7]. From a functional perspective, understanding how GPIAPs are distributed in cell membranes has implications for the large variety of proteins with GPI anchors, and for many of the putative roles ascribed to the GPI anchor [8]. The relationship between the functional and structural (or morphological) organization of GPIAPs in cellular membranes will be explored in this chapter.

GPI-Anchor Function and the ‘‘Raft’’ Hypothesis GPIAPs, in principle, resemble glycolipids with abnormally large head groups, and therefore they should have properties similar to cellular glycolipids. GPI PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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anchors provide a relatively stable membrane anchor for proteins at the extracytoplasmic leaflet of the plasma membrane. However, over several hours or days, GPIAPs associated with cell membranes may redistribute to neighboring membranes. Therefore, the association of GPI anchors with cellular membranes may be weak in comparison with the strength of transmembrane protein-based anchors. This property may have clinical potential for ‘‘painting’’ cells with antigens of interest in vivo [9, 10]. The GPI anchor acts as an apical targeting signal for proteins in many epithelial cell types [11–13]. GPI anchors do not extend across the bilayer, and for sorting to be realized, the anchor may associate with transmembrane proteins which are themselves distributed in the plane of the membrane, permitting segregation in the plane of the bilayer into specialized domains. GPI anchoring has also been shown to be important for the intracellular signaling capacity of several lymphocyte proteins; in most cases, cross-linking of the protein is a prerequisite for signaling [14, 15]. The intracellular signaling capacity of GPIAPs requires a mechanism for transmitting a signal across the bilayer, possibly by association with a transmembrane linker or by virtue of association with specialized microdomains. Simons and co-workers have put forward a unifying hypothesis, the ‘‘raft hypothesis,’’ that provides a framework to understand these functions [16]; the glycosphingolipid and cholesterol-enriched areas of membranes or ‘‘rafts’’ recruit a specific set of membrane proteins (including GPIAPs) and exclude others, thereby sorting membrane proteins or concentrating signaling molecules where they may interact more efficiently with their effectors. Because domains and rafts have been used to describe a variety of structural features in a cellular context, it is useful to define these terms in context. In cells, there are domains that constitute morphological specialization of the plasma membrane, such as the apical and basolateral domains in polarized epithelia or the axonal and dendritic surfaces of neurons. These domains arise due to active processes that are mediated and maintained by complex cellular machinery. Another type of domain or raft arises due to lateral heterogeneities in relatively undifferentiated areas of the plasma membrane (such as areas devoid of morphologically defined specialization) in polarized and nonpolarized cells. It is these rafts that are thought to play a role in the biological manifestation of GPIanchor function.

Lateral Heterogeneity in Cell Membranes Structurally, GPIAPs are (glyco)lipids with unusually ‘‘large’’ head-(protein) groups, and their organization in the membrane may be understood in the context of the distribution of other membrane components, especially glycolipids. Our view of the structure and organization of the membrane of cells has undergone major revisions from the time of the fluid mosaic model proposed by Singer and Nicolson [17]. The membrane is no longer considered as a ‘‘two-dimensional oriented solution of integral proteins . . . in a viscous phospholipid bilayer,’’ but rather both protein and lipid components of the bilayer are arranged nonrandomly in lateral and vertical directions. Asymmetries arise due to protein– protein, lipid–protein, or lipid–lipid interactions [18, 19], or as in the case of

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transbilayer asymmetry of membrane lipids, to active processes such as the asymmetric synthesis and flip–flop of some lipid classes [20]. Before examining the evidence for the presence of lateral segregation or rafts in cell membranes it is useful to survey biophysical studies in different artificial membrane systems. A clear indication of lateral heterogeneity is the coexistence of phase-separated regions in the membrane bilayer, validated by both theoretical considerations and experimental observations (reviewed in [21, 22]). Segregated domains in artificial bilayers may arise due to many different types of interactions between membrane components [18, 19, 21, 23]. In a bilayer of lipids with mismatched acyl chains, small differences in interaction energies will lead to the segregation of liquid-ordered regions (rigid domains) consisting of long saturated acyl chain lipids. In contrast, the shorter acyl chain or unsaturated acyl chain lipids prefer the liquid-disordered, less rigid domains [22]. Preferential lateral interactions between hydrophilic head groups and strong lateral interactions between cholesterol and sphingolipids are also obvious mechanisms for lateral segregation [21]. Cholesterol (and possibly other sterols) may play a major role in modulating the size and extent of these domains, due to their planar ‘‘rigid’’ structure which may potentiate the organization of liquid-ordered domains in membranes [22, 24, 25]. Although, studies in artificial systems do not reveal the architecture of rafts in the plasma membrane, they provide guidelines for investigating this phenomenon in cells.

Detergent-Resistant Membranes and Their Connection with Rafts An important early clue to the function of rafts was the observation that the apically sorted viral protein hemaglutinin (HA) was insoluble in nonionic detergents, both at the Golgi and at the plasma membrane [26]; apically targeted GPIAPs were also insoluble in cold detergents at the apical surface of epithelial cells and at the Golgi en route to the apical surface [27]. At about the same time, GPIAPs were shown to be distributed in clusters, concentrated over caveolae [28, 29]. Caveolae are small flask-shaped cell surface invaginations, covered with a striated proteinaceous coat, the main component of which is the cholesterolbinding protein, caveolin [29, 30]. In addition, GPIAPs were also found in lowdensity, detergent-resistant membrane fractions in association with caveolin [31]. From these data, an association of GPIAPs was inferred with caveolae in specific detergent-resistant domains or in rafts in the membrane, which are potentially involved in polarized sorting [27, 32]. Much of the proof for the existence of domains or rafts in cells rests on the same basic result that specific membrane proteins and lipids, including not only GPIAPs but also lipid-linked nonreceptor tyrosine kinases, (glyco)sphingolipids, caveolin, and cholesterol form low-density detergent-insoluble complexes in cold Triton X-100 extracts of cells [16, 32, 33]. However, copurification with such detergent-insoluble material does not constitute proof of preexisting domains in cells. The detergent itself may cause a coalescence or removal of components; therefore, these structures are likely to be different from the domains or rafts hypothesized to be involved in functional processes inside cells [34–36]. This is

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an obvious but poorly appreciated point, and several researchers at different times have equated detergent-resistant complexes with caveolae, signaling complexes consisting of nonreceptor tyrosine kinases, and sorting domains (see [32, 36, 37]).

Morphological Correlates of Detergent-Resistant Membranes Analyses of the cell surface distribution of different GPIAPs using fluorescence microscopy and immune-electron microscopy have shown that these proteins are not concentrated in caveolae or in small ‘‘puncta’’ or clusters at all times (Fig. 10-1A and C); rather, they are enriched in these structures only after crosslinking with polyclonal secondary antibodies [38–40]. Careful electron microscopic (EM) examinations have shown that, to the level of sensitivity of the technique, GPIAPs are neither concentrated in nor excluded from caveolae or clathrin-coated pits; cross-linking with antibodies redistributes these proteins into clusters which are enriched in morphologically identified caveolae [35, 38–40] (see also Fig. 10-1C). One study has suggested that there may be small clusters of GPI-anchored folate receptors detectable by EM analysis, under conditions which do not perturb the aggregation state of these molecules [41]. In general, however, multimerization of GPIAPs, apparently causes sequestration in caveolae, but in the absence of clustering agents they are uniformly distributed over the plasma membrane. After detergent extraction, the remaining large sheets of fenestrated membrane are devoid of transmembrane proteins and cytosolic components, and GPIAPs and other detergent-insoluble species appear to be associated with these sheets (Fig. 10-1B and D) [35]. At the EM level, these membrane sheets appear to be formed of a bilayer, and they exhibit an enhanced degree of clustering of the GPIAPs (Fig. 10-1D) [35].

‘‘Signals’’ for Detergent Insolubility The insolubility of a large fraction of cellular GPIAPs, along with the lack of enrichment in the relatively small punctate areas occupied by caveolae after detergent treatment, indicates that these proteins are detergent-insoluble independent of their association with caveolae. The lack of association with caveolin/ VIP-21 is consistent with studies in lymphocytes, where GPI-anchored Thy-1 and the ganglioside GM1 were detergent insoluble and almost quantitatively formed ‘‘low-density complexes’’—even though the cells lack detectable caveolin/ VIP-21 or caveolae [42]! Such observations, obviously, raise questions about how GPIAPs, other lipid-linked molecules, and transmembrane components become enriched in the insoluble residue from detergent-extracted cells. Cytoplasmically oriented proteins such as nonreceptor protein tyrosine kinases (NPTKs) and heterotrimeric G proteins also have been found in detergent insoluble complexes [31, 43–47]. These proteins share a familiar posttranslational modification: closely juxtaposed saturated fatty acyl chains. In most cases, these proteins are N-myristoylated on the N-terminal glycine and palmitoylated at a cysteine residue near the N-terminal glycine. These data favor the hypothesis that association of the detergent-insoluble membranes requires closely juxtaposed


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Figure 10-1 Surface distribution of GPI-anchored proteins. Folate receptor-expressing CHO cells were incubated with a fluorescent analogue of folic acid, PLF, and either treated with buffer without Triton X-100 (A) or with 1% Triton X-100 (B) on ice for 20 min [35]. The cells were then imaged immediately as described in [35]. Note uniform distribution of folate receptor at the cell surface prior to Triton X-100 treatment and fenestrated appearance of the cell membrane after Triton X-100 treatment. Greater than 90% of the folate receptor remains with the detergentinsoluble pellet under these conditions, whereas more than 95% of a transmembrane-anchored protein, the transferrin receptor, is solubilized [35]. Bar, 10 애m. Decay-accelerating factor (DAF) expressing CHO cells [35] were incubated with mAbs to DAF (1A10) and either treated with buffer without Triton X-100 (C) or with 1% Triton X-100 (D) on ice for 20 min. The cells were then fixed with 3% paraformaldehyde and 0.5% glutaraldehyde, labeled with goldconjugated antibodies (10 nm) to detect the mAb, and processed for electron microscopy as described in [35]. Prior to Triton X-100 treatment, the GPIAP appears randomly distributed in the plasma membrane, neither excluded from coated pits nor from caveolae at the cell surface. After detergent treatment, the membrane appears to retain its bilayer nature but is present as fragments in the section shown, consistent with the fenestrated appearance observed in panel B. The gold particles appear to be more clustered after Triton X-100 treatment compared to the untreated condition. For a quantitative analysis of the above, see [35]. Bar, 100 nm.

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saturated fatty acyl chains and the presence in the same bilayer of the various saturated acyl/alkyl chain-containing proteins and lipids [48, 50]. In confirmation of this hypothesis, the association of two NPTKs, p56 lck and p59 fyn with detergentinsoluble complexes, was found to be due to the N-terminal myristoylation at the Glycine residue and palmitoylation at the Cys 3 residue [48]. Further proof came from recent work from Arni and co-workers, who showed that the association of GAP-43 with detergent-resistant membranes required two palmitoylated cysteine residues, Cys3 and Cys 4, and this signal for detergent insolubility is transferable to the N terminus of 웁-galactosidase [49]. Cytoplasmic lipid-modified proteins also share the property of detergent insolubility and association with low-density complexes. However, it should be noted that at least in the case of the GPIAPs, this association does not reflect an initial concentration in any specialized structures [35].

The Physicochemical Basis for Detergent Insolubility The basis of detergent insolubility of GPIAPs has received considerable attention (reviewed in [50, 51]). The insolubility of GPIAPs and membrane lipids in Triton X-100 can be reconstituted in the absence of any special structures or protein(s) [52]. The main requirement for detergent insolubility is the presence of a significant mole fraction of high-melting temperature lipids such as saturated acyl chain-containing phospholipids. Decreasing the concentration of cholesterol or neutral glycolipids negatively influences the extent of insolubility. GPIAPs incorporated into such liposomes are insoluble in cold nonionic detergents and can be solubilized by increasing temperature. In contrast, the lipid component is equally insoluble at high and low temperatures, suggesting that the GPIAPs are only loosely associated with the detergent-resistant membranes. Furthermore, detergent insolubility of GPIAPs requires the presence of the appropriate lipids in the same bilayer as these proteins [52]: insolubility depends mainly on the acyl or alkyl chain composition of the membrane lipids, cholesterol, or neutral glycolipid content, and probably the degree of saturation of the acyl or alkyl moiety of the GPI anchor. In most GPIAPs, saturated alkyl/acyl chains appear to be the predominant components of the lipid portion of GPI anchors [7] which contribute to their detergent insolubility. The requirements for phase segregation of lipids in the fluid phase into a disordered phase (liquid-disordered) and a more ordered phase (liquid-ordered) wherein acyl chains of phospholipids have properties intermediate between gel and fluid phases and for generating detergent-insoluble complexes appear similar. Cholesterol and sphingolipids are not essential for insolubility in artificial liposomes but potentiate the Triton X-100-insolubility of the incorporated GPIAPs [25, 53]. Modulating cholesterol levels in vivo by treatment with agents that inhibit cholesterol synthesis (like compactin [54, 55]) or alter its distribution (such as saponin [25, 53, 56]) make GPIAPs more soluble in detergent by reducing the available cholesterol in membranes. In a sphingolipid, deficient cell line insolubility of GPIAPs is considerably reduced as sphingolipid levels are lowered; insolubility of the GPIAP could be restored to control levels by metabolic compensation with exogenously added sphingolipids [54]. These data are consistent with the hypothesis that detergent-insoluble domains require cholesterol and sphingolipids. However, the composition of the detergent-resistant membranes has not been de-


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termined. As mentioned above, cholesterol and sphingolipids enhance the detergent insolubility of GPIAPs by promoting the formation of ordered membrane domains, which are likely to be insoluble in Triton X-100 [25, 52, 53]. These domains are relatively stable in fluorescence quenching experiments [25]. GPIAPs can be isolated from both cells and sphingolipid and cholesterolrich liposomes (SCRLs) in association with detergent-insoluble membranes. Detergent insolubility of lipids was characteristic of phases in which the lipid acyl chains are predominantly in the liquid-ordered phase as shown in studies of Brown and co-workers [22, 53]; GPIAPs become insoluble because they associate with cholesterol and sphingolipid-rich lipid domains with properties similar to the liquid-ordered phase. Saponin extraction had similar effects on one GPIAP, alkaline phosphatase, that had been incorporated into SCRLs. Direct interactions between GPIAPs and cholesterol were not required, because the protein was also detergent insoluble in cholesterol-free liposomes containing lipids in an ordered phase. These studies strongly suggest that the liquid-ordered domain model can explain the generation of detergent-resistant membranes in liposomes and cells [53]. In conjunction with other results [25], the available data suggest that cholesterol merely acts to potentiate the degree of insolubility correlated to its ability to influence liquid-ordered phase formation in membranes. In summary association with the detergent-insoluble membranes requires closely juxtaposed saturated fatty acyl chains and the presence in the same bilayer of the various saturated acyl/alkyl chain-containing proteins and lipids that can associate with or promote the formation of liquid-ordered phases. However, at present, it is unclear whether the biochemically defined detergent-resistant membranes and the functional entities termed rafts are related [22, 36, 50].

Evidence for Rafts in Living Cell Membranes As mentioned at the outset, the most compelling evidence for domains in the undifferentiated plasma membrane of cells is functional. While several investigators have looked for domains by techniques capable of detecting lateral segregation over different time and length scales, convincing evidence for rafts in living cell membranes has been elusive until very recently [57]. Rafts in living cell membranes are likely to differ in size and structure from the phase-segregated regions or ‘‘self-organized’’ lipidic domains observed in artificial systems, but similar physical principles must apply to both [19, 22, 57]. However, many biophysical measurements, including differential scanning calorimetry, spectroscopy, and NMR, have been inconclusive in detecting selforganized domains in living cells, probably due to the presence of a relatively large number of lipid species and the presence of proteins, some of which have specific lipid annuli associated with their hydrophobic transmembrane domains. Studies of membrane protein diffusion using microscopic techniques such as fluorescence recovery after photobleaching (FRAP), single particle tracking, and optical laser traps at the cell surface have shown that proteins do not undergo the exclusively unrestricted lateral diffusion characteristic of a relatively dense, two-dimensional fluid [58]. Proteins appear to have many modes of motion at the cell surface: Brownian diffusion, diffusion which appears to take place in transiently confined regions (corralled diffusion), and modes of motion which

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do not have characteristics of Brownian motion (anomalous diffusion) [59]. A considerable fraction of membrane protein is immobile over long time scales as measured by FRAP experiments and single-particle tracking analyses. In addition, they are transiently confined for average times of 6–8 s to small domains (200–400 nm in diameter) in relatively undifferentiated areas of the membrane [60]. However, the size and duration of transient confinement do not appear to be dependent on the nature of the membrane anchor of the protein: GPIanchored or transmembrane-anchored neuronal cell adhesion molecules, and the ganglioside GM1 exhibit similar diffusive motions at the surface of plasma membrane [60, 61]. These observations may, instead, reflect considerable lateral heterogeneity in the membrane at the nanometer scale [59, 62]. There appears to be a consensus picture emerging for the underlying bases for these lateral constraints on ‘‘free diffusion’’: proteins may be confined in membranes by the apposed cytoskeleton, or by lipid microphases such as liquid-ordered phases of the scale of transient confinement zones in the membrane. Collectively, the observations made above suggest that there are domains or corrals that constrain lipid-linked molecules in the plane of the membrane, at least at the extracytoplasmic leaflet of the bilayer. The existence of rafts, however, requires the lateral segregation of proteins in membranes: it is this concentration that has been difficult to establish by conventional fluorescence microscopy and electron microscopic methods. Failure to obtain evidence for this defining property of rafts may be due to their dynamic state on the usual time scales of imaging methods and the limited resolution of current microscopic techniques. Fluorescence microscopy is limited by the wavelength of the emitted light (⬎300 nm) [63], and electron microscopic techniques are too insensitive to detect small aggregates of lipidic species [64] (but see [41]). Therefore, domains or rafts must be small (much less than 300 nm in size), dynamic, and contain only a few protein species [19]. Recently, two studies [65, 66] using very different techniques have provided the first consistent evidence for an unusually high density of GPIAPs in cell membranes independent of their overall concentration in the cell, thereby fulfilling the criterion of lateral segregation in the membrane. In the first, Friedrichson and Kurzchalia utilized classical chemical cross-linking techniques to show that GPIAPs are present as molecular aggregates consisting of about 15 molecules [65]. Chemical cross-linking by a cell impermeant cross-linker, bis(sulfosuccinimidyl)suberate (BS3), resulted in a series of cross-linked multimers of GPI-anchored isoforms of the normally secreted and monomeric human growth hormone receptor and the naturally occurring GPI-anchored folate receptor. The extent of crosslinking did not correlate with expression levels of the GPIAPs in membranes, but was found to be critically dependent on the presence of a GPI anchor; transmembrane-anchored isoforms did not exhibit similar cross-linked species. Furthermore, cholesterol depletion of membranes reduced the extent of crosslinked species, while, detergent treatment dramatically increased the extent of cross-linking. In the second study, Varma and Mayor have used a new fluorescence resonance energy transfer (FRET) methodology to ascertain, at a nanometer scale, the organization of GPIAPs in living cell membranes [66]. This method is based on a property of FRET which results in a loss in polarization of the emission from a fluorophore that has been excited by polarized light [67]. This loss in


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polarization (or anisotropy) is mainly the result of the relatively large allowed spread in the orientations of the acceptor fluorophores and the random orientational spread of the acceptor molecules. The extent of depolarization depends on the usual sixth power of distance between transferring species, and may be calibrated like conventional FRET in all other respects. The loss of fluorescence anisotropy provides a very sensitive method to assay for FRET even between like fluorophores [68], because anisotropy is a physical quantity which can be measured with great precision. Varma and Mayor compared the organization of GPIAPs to a transmembrane protein in Chinese hamster ovary (CHO) cells transfected with a GPIanchored folate receptor (FR-GPI) or a chimeric transmembrane-anchored folate receptor (FR-TM). The folate-binding domain was used as a reporter molecule because it is a monomeric protein and can be exogenously labeled by binding monovalently to a fluorescent folic acid analog, N움-pteroyl-N␧-(4⬘-fluoresceinthiocarbamoyl)-L-lysine (PLF) [35]. Consistent with the data from fluorescence

Figure 10-2 Possible outcomes of the fluorescence depolarization experiment for different arrangements of fluorescently labeled GPI-anchored proteins in a microscope pixel. If GPI-anchored proteins are present as uniform random distributions in membranes (A), the average distance between individual protein molecules on a two-dimensional surface increases by a factor of two upon a fourfold decrease in fluorophore density (or number, n). Because the fluorescence intensity measured by a pixel of a CCD camera is representative of the concentration of the fluorophores on the cell surface imaged by that pixel, the polarization (or anisotropy, r) values of two pixels differing in fluorescence intensity will be different if the fluorophores are within energy transfer distances of each other. If GPI-anchored proteins are arranged in subpixel-sized domains or rafts at the cell surface (B), the local fluorophore density would be proportional to the number of such domains in a given pixel, and the distance between fluorophores would be independent of the fluorophore density. If these domains are submicron sized, the anisotropy values of two pixels differing in fluorescence intensity will be independent of fluorophore intensity. (Reprinted with permission from Nature [66], copyright 1998 Macmillan Magazines Limited.)

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and electron microscopy (Fig 10-1A and C), only two possible arrangements of fluorophores are possible as visualized in a microscope pixel focused at the cell surface (Fig. 10-2). These arrangements will have a predictable relationship of anisotropy and pixel fluorescence intensity according to their known geometry. Fluorescence anisotropy values at the surface of FR-GPI-expressing cells were constant over the entire fluorescence intensity range (Fig. 10-3a and b, see color plate), whereas anisotropy values were inversely dependent on fluorescence intensity for the transmembrane-linked protein, FR-TM (Fig. 10-3c and d). The constant anisotropy value for different fluorescence intensities seen for FR-GPI suggested that there was no energy transfer taking place between the FR-GPI species, or alternatively, GPIAPs were organized in small, subpixel resolution domains at the cell surface such that measurable energy transfer occurred between fluorophores (see Fig. 10-2B). Of these possibilities anisotropy was found to be relatively depolarized due to energy transfer, because methods that reduced local fluorophore density, dilution by folic acid competitor or photobleaching of the fluorophore at the surface of cells, resulted in increased anisotropy [66]. Cholesterol also altered the arrangement of the GPIAPs at the cell surface. Varma and Mayor have recently observed that the presence of a different GPIAP [decay accelerating factor (DAF)] in the membrane, increases the anisotropy of PLF-labeled FR-GPI in a systematic fashion; higher values of anisotropy for PLF-labeled FR-GPI correlate with higher DAF expression levels relative to FR-GPI levels in the same cell (Varma and Mayor, unpublished observations). These results provide evidence for the presence of multiple GPIAPs in these domains. However, Kennworthy and Edidin, using more conventional FRET methodology, have come to very different conclusions. They used energy transfer between Cy3 and Cy5-fluorophore-labeled antibodies against 5⬘-nucleotidase (5⬘NT) to study the organization of GPI-anchored 5⬘NT in transfected epithelial cells [69]. The extent of energy transfer correlated with increasing protein density, inconsistent with a model in which 5⬘NT is constitutively clustered in the membrane. Differences between these two studies cannot be reconciled at present. One possible explanation may be that in the epithelial cells used by Kenworthy and Edidin, a significant preexisting pool of endogenous GPIAPs ‘‘dilutes’’ exogenously transfected 5⬘NT; the extent of energy transfer would relate to higher expression levels of 5⬘NT. Studies using fundamentally different methodologies provide direct evidence for the lateral segregation of GPIAPs in rafts or in domains in living cell membranes. These experiments, however, do not provide information regarding the structure, size, dynamics, and stability of the rafts in question, but, at best, place an upper bound on any of these parameters.

Involvement of Rafts in Biosynthetic and Endocytic Sorting Sorting of proteins to different routes during biosynthetic transport is mediated by specific protein sequence or glycosylation-based motifs (such as the targeting of lysosomal membrane proteins to lysosomes from the trans-Golgi network [70], or the delivery of specific sets of proteins to the basolateral and apical domains of


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epithelial cells) [36, 71]. The realization that lipid-based motifs are also involved in sorting of membrane components comes from the observation that glycosphingolipids and GPIAPs are preferentially directed to apical domains in epithelial cells [13, 72]. That GPIAPs, glycosphingolipids, and cholesterol are found in detergentresistant complexes during biosynthetic transport to the apical membrane, along with a canonical apical protein, the HA protein of influenza virus, provided support for the involvement of membrane rafts or detergent-resistant membranes in sorting [16, 27, 73, 74]. However, there are several reasons to question the conclusion that the detergent-insoluble complex is a ‘‘sorting platform.’’ Sphingomyelin, although a major component of the detergent-resistant lipids, is not enriched at the apical surface [75]. Replenishment of lipids incapable of being incorporated into detergent-resistant membranes rescues the impaired sorting of GPIAPs in sphingolipid-depleted cells [76, 77]. Furthermore, a moderate depletion of cholesterol results in the loss of the ability of GPIAPs to associate with detergent-insoluble complexes, while they are still sorted to the apical domain [55]. Severe depletion of cholesterol does result in the loss of polarized delivery of HA in MDCK cells [78]. The capacities of proteins to associate with detergent-resistant membranes and to sort to the apical surface appear to be correlated and are resident in the transmembrane region of the protein. Mutations that alter detergent insolubility, in general, also affect the ability of the mutant HA to apically sort [79]. The involvement of the GPI anchor as a general apical targeting signal has also been challenged. In Fisher rat thyroid cells, which are epithelial cells that do not express the caveolin protein, GPIAPs are targeted to the basolateral surface, and the ectopic expression of caveolin does not reverse this phenotype [80–83]. A non-GPI-anchored truncation mutant of a normally GPIAP, placental alkaline phosphatase, retained its apically polarized delivery [84], suggesting that the GPI anchor is not essential for apical delivery. While these observations do not exclude involvement of rafts in the sorting process, they reflect the complexity of biosynthetic sorting in the TGN and imply a hierarchy of signals that dictate sorting in polarized epithelia. Several lines of evidence suggest that rafts are involved in sorting in the endosome, but the endocytic route of GPIAP internalization also has been a subject of much controversy. Although GPIAPs are distributed throughout the plasma membrane, they may be internalized by specific routes in some cells, especially, when they are artificially cross-linked [85]. In general, glycolipidcontaining areas of plasma membrane and GPIAPs (including the folate receptor) are internalized in noncoated vesicles and in the clathrin-coated pathway at rates corresponding to overall membrane internalization [86–88]. GPIAPs are internalized into true endosomes which contain recycling transferrin receptors and are quantitatively recycled back to the cell surface [88]. However, as compared to recycling receptors and most membrane components, GPIAPs are extensively retained in endosomes [88, 89]. Retention depends on the lipid levels in cellular membranes: cholesterol and sphingolipid depletion appear to relieve retention relative to other recycling receptors [77, 88]. In functional terms, lipiddependent retention in endosomes exposes GPIAPs to the acidic milieu of both sorting and recycling endosomes for longer times than recycling receptors with conventional transmembrane anchors. This behavior explains for the GPI-anchor

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and cholesterol and sphingolipid-dependent uptake of folates by the folate receptor [90, 91], and the formation of infectious prion protein from the cellular scrapie form [92–94]. If infectious forms of the scrapie protein were present in endosomes, the cholesterol-dependent retention mechanism would facilitate an efficient conversion of the native scrapie protein into prions, possibly in distinct membrane domains or aggregates [94]. Recent data supportive of the role of specific lipid domains (rigid- or liquidordered domains) segregating from other types of membrane (liquid-disordered or fluid areas) come from studies on membrane organization and the trafficking properties of the lipophilic dialkylcarbocyanines, the rigid-domain-preferring DiIC16 and the fluid-domain-preferring DiIC12 in mammalian cells [95–97]. The two kinds of lipid analogues are sorted from each other at the surface of cells [95, 96] and in endosomes [97], consistent with the hypothesis that the rigiddomain molecules may segregate into specialized cholesterol-rich rafts in endosomes and at the cell surface. In spite of the circumstantial evidence described above, the direct involvement of rafts in biosynthetic sorting in the TGN, and during sorting in endosomes, remains to be demonstrated [36, 50].

Rafts and Signaling by GPIAPs Many signaling proteins expressed on lymphocytes are GPIAPs, including Thy1, Qa-2, FcR웂-III, and LFA-3 [73] (see Chapter 8 by Schofield). A variety of intracellular responses may be elicited by cross-linking different GPIAPs on T lymphocytes [15]. These events include a rise in intracellular calcium, protein tyrosine phosphorylation, interleukin-2 production, and in some cases proliferation. An early event in the initiation of GPIAP signaling appears to be the activation of lipid-linked nonreceptor tyrosine kinases (NPTKs) on the cytoplasmic leaflet of the plasma membrane [33]. Due to the lack of any membrane extensions for both the classes of proteins, the mechanism by which signal transduction can occur remains obscure. The idea that rafts in cell membranes may serve to organize GPIAPs on the outer leaflet and lipid-linked nonreceptor tyrosine kinases on the inner leaflet is an attractive hypothesis, but requires that rafts exist in both leaflets of the plasma membrane. As yet it is unclear whether domains at the cell surface can induce domains at the cytoplasmic leaflet, or vice versa, or if rafts are present at all at the cytoplasmic face of the plasma membrane. Recent experiments suggest that after cross-linking, GPIAPs patches partially colocalize with c-fyn in T lymphocytes [98]. Further studies will be required to distinguish whether cross-linked GPIAPs could cause the formation of membrane domains which assemble signaling molecules, or if GPIAPs and cytoplasmic signaling molecules exist in complexes in microdomains which are unresolvable at the resolution of current localization techniques. In the absence of direct evidence in living cells, experiments in artificial membranes suggest possible mechanisms whereby cholesterol may link domains across the bilayer, because cholesterol has the potential to form end-to-end dimers in the membrane [99, 100]. Signal transduction by GPIAP in T lymphocytes is impaired by lowering cellular cholesterol content or by feeding cells


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liquid-ordered-phase disordering lipids such as polyunsaturated fatty acids [101, 102]. Consequently, the establishment of raft association of the signaling components, namely lipid-linked NPTKs, at the inner leaflet of the plasma membrane is central to understanding signaling by these molecules. In a recent study, Illangumaran and co-workers [103] have shown that NPTKs are present in high (heavy) and low density (light) plasma membrane fractions in distinct forms. The light membranes are enriched in GPI, whereas the heavy membranes contain the bulk of the plasma membrane lipid and protein, and they have several other components associated with them. These investigators propose that the two fractions represent distinct membrane domains, the light membrane consisting of core GPIAP and sphingolipid-enriched membranes while the heavy membranes consist of the GPIAPs and other transmembrane components that may laterally associate with these domains. Different membrane environments are inferred because the GPIAP-enriched ‘‘light’’ fractions show differential activation of NPTK activity upon detergent extraction or are made permeable with specific cholesterol sequestration agents. Thus far, it is not clear that NPTKs are associated with any form of lateral heterogeneity, which has an identity outside the detergent-insoluble complex paradigm. In a recent study, Stauffer and Mayer have shown that green fluorescence protein (GFP) tagged Src-homology domains, which are present in all the NPTKs, are recruited to GM1-enriched domains after IgE-receptor triggering [104]. In separate studies, functional signaling mediated by T cell receptor activation during antigen presentation also suggests the involvement of rafts [105]. These observations together strengthen the raft hypothesis, but conclusive proof of the preexistence of rafts of NPTKs at the cytosplasmic leaflet of the cell membrane is lacking.

In Conclusion Understanding of GPIAP function clearly points to the association of these proteins with lateral segregations or rafts that participate in cell membrane associated function. Recent evidence for the existence of such structures at the cell surface provide some evidence for the viability of the raft hypothesis, but there are a host of unanswered questions regarding the physical nature of these entities and their mechanism of participation in specific cellular functions. It is important to appreciate that the understanding of rafts as functional units inside the cell will depend primarily on context. Rafts are involved in sorting a host of proteins (especially GPIAPs) at the TGN and in endosomes; at the plasma membrane rafts are involved in transducing signals across the bilayer. At the present time, it is unclear whether similar physical structures are involved in both functions. These problems and new methodologies to address them will centrally occupy research on GPIAP function for some time.

References 1. Ferguson, M. A. J., and Williams, A. F. (1988). Cell-surface anchoring of proteins via glycosyl phosphatidylinositol structures. Ann. Rev. Biochem. 57, 285–320.

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11 Structure and Function of GPI-Specific Phospholipases Martin G. Low Department of Physiology and Cellular Biophysics, Columbia University, New York, New York 10032

Introduction: GPI Anchors and Phospholipases As indicated in the preceding chapters, glycosylphosphatidylinositol (GPI) anchors are widely distributed among cell surface proteins in eukaryotic organisms. The GPI anchor has an essential structural role in the attachment of proteins to the membrane, but many questions remain about its functions, and a decade after its discovery, the utility of the GPI anchor compared to the conventional transmembrane polypeptide remains unclear. A unique feature of GPI anchors is their ability to be cleaved by specific phospholipases resulting in the release of the protein from the membrane. This property has proven of great practical value and played an important role in the initial discovery and characterization of GPI-anchored proteins. In fact, the history of these two biochemical entities, the GPI anchors and their phospholipases, is so intertwined that it is difficult to perceive which was discovered first [1, 2]. Three distinct types of phospholipase are capable of hydrolyzing the phosphodiester bond linking the lipid moiety and the inositol residue in GPIs. This chapter describes these ‘‘GPI-specific’’ phospholipases, with particular emphasis on the mammalian GPI-specific phospholipase D(GPI-PLD). The biochemical properties of these enzymes are compared in Table 11-1. Also included in the table for comparison is the mammalian phosphoinositide-specific phospholipase C family; these enzymes can hydrolyze phosphatidylinositol and polyphosphoinositides but appear unable to degrade GPIs. PNH and the GPI-Linked Proteins Copyright © 2000 by Academic Press. All rights of reproduction in any form reserved.

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TABLE 11-1 Comparison of Phospholipases Which Hydrolyze Inositol Phospholipids Phosphoinositide PLC

PI-PLC

GPI-PLC

GPI-PLD

Major source

Mammalian cells

Some bacteria

T. brucei

Molecular mass Substrate specificity

앑85–155 kDa PI, PI 4-P, or PI 4,5-P2 Ca 2⫹

앑35 kDa PI or GPI

앑37–40 kDa PI or GPI

Mammalian plasma Some cell types 앑100–120 kDa GPI or GPI(acyl)

None

None

Ca2⫹, Zn 2⫹

Inositol 1,2-cyclic P and inositol 1-P a

Inositol 1,2-cyclic P a or glycan inositol 1,2-cyclic P b 1,2-Diacylglycerol d Secreted

Inositol 1,2-cyclic P a or glycan inositol 1,2-cyclic P b 1,2-Diacylglycerol d Intracellular membranes

Glycan inositol b or glycan inositol(acyl)c

Metal ion involvement Polar product(s) a,b

Lipid product Subcellar location

1,2-Diacylglycerol Cytoplasmic

Phosphatidic acid Secreted

With a PI, b GPI, or c GPI(acyl) as substrate; d lipid product can also be 1-alkyl, 2-acyl glycerol, etc., depending on the lipid moiety in the intact GPI anchor. P, phosphate.

Phosphatidylinositol-Specific Phospholipase C Phosphatidylinositol-specific phospholipase C (PI-PLC) is secreted into the media by several different gram-positive bacterial species, including Staphylococcus aureus, Bacillus cereus, Bacillus thuringiensis, Clostridium novyi, and Listeria monocytogenes. Strictly speaking, these enzymes are not GPI specific because they will cleave phosphatidylinositol as well as GPI. However, they do not hydrolyze the polyphosphoinositides (phosphatidylinositol 4-phosphate and phosphatidylinositol 4,5-bisphosphate) or major membrane phospholipids such as phosphatidylcholine, and phosphatidylethanolamine. Their relatively narrow substrate specificity combined with an ability to act on membranes and intact cells has made the bacterial PI-PLCs very useful tools, not only to identify novel GPI-anchored proteins but also for functional studies [3, 4]. Staphylococcus aureus PI-PLC, although widely used in early studies, suffers from two practical disadvantages: relatively low specific activity compared to the Bacillus enzymes and strong inhibition by the salt concentrations present in mammalian cell culture media. Similarly, the PI-PLC from L. monocytogenes is reported to have relatively poor activity toward GPI-anchored proteins substrates compared to PI [5]. Consequently the two Bacillus enzymes (which are essentially identical in their sequences and biochemical properties) are currently favored for identifying and characterizing free GPIs and GPI-anchored proteins [3, 4]. The crystal structures of the mammalian and two bacterial PI-PLCs reveal a high degree of structural conservation in the substrate binding site, consistent with their distinct but overlapping substrate specificity [6–8]. The structural comparison also suggests that both types of PI-PLC evolved from a common ancestor and then diverged for functional reasons [9]. The bacterial enzyme has retained its size, whereas the mammalian PI-PLC has acquired additional domains allowing its activity to be regulated, with great precision, by the intracellular

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environment. Consistent with this hypothesis is the recent discovery of two distinct PI-PLC activities in Streptomyces lividans [10]. Although both of the Streptomyces enzymes will hydrolyze PI, only one of them is capable of hydrolyzing GPI; a comparison of their amino acid sequences, reaction mechanism, and Ca2⫹ requirements suggests that one is very similar to the bacterial PI-PLC, whereas the other is more closely related to the mammalian enzyme [10]. An important (and useful) feature of the catalytic mechanism utilized by bacterial PI-PLC is that cleavage of the inositol phosphate headgroup from the diacylglycerol is not a simple hydrolytic reaction [11]; rather, an intramolecular phosphotransfer first produces a relatively stable inositol 1,2-cyclic phosphate intermediate (Fig. 11-1). This reaction is then followed by a slow hydrolysis of the cyclic phosphate to produce the inositol 1-phosphate [9]. The second reaction is so sluggish that, under normal conditions (incubation periods just sufficient to get complete cleavage of PI), it is not detectable by standard techniques. By contrast, the mammalian PI-PLCs, for which the second reaction is quite rapid, produce approximately equal amounts of inositol 1,2-cyclic phosphate and inositol 1-phosphate. This rather esoteric difference between the two types of enzyme is of great practical value when GPI-anchored proteins are the substrates. The inositol 1,2-cyclic phosphate moiety, which is exposed when GPIs are cleaved by bacterial PI-PLC, is immunogenic and as a consequence is usually the dominant

Figure 11-1 Cleavage of GPI anchors by phospholipases C and D. GPI anchors can be cleaved by either a phospholipase C or D type mechanism. The initial reaction of phospholipase C mediated cleavage involves an intramolecular phosphotransfer to release a product terminating with an inositol 1,2-cyclic phosphate moiety. The GPIs shown here contain a diacylglycerol, but the reactions are essentially the same for GPIs with different lipid moieties. For clarity, the acyl groups have been truncated (dotted lines). Ins, myo-inositol.

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epitope in CRD (or ‘‘cross-reacting determinant’’) antisera [12–16]. CRD antisera have been extensively used for the analysis of GPIs cleaved by PI-PLC by immunoblotting techniques (see later and Table 11-1). However, it should be pointed out that excessive amounts of PI-PLC or prolonged incubation periods will eventually hydrolyze the 1,2-cyclic phosphate and destroy the CRD. The function of the bacterial PI-PLCs is unknown. Most bacterial species do not contain inositol phospholipids, so it is unlikely that PI-PLC is required for PI metabolism or for the salvage of inositol from their natural environment. Improbable also is a role for PI-PLC in the metabolism of fatty acids, glycerol, and phosphate; much larger amounts of these nutrients would be available from more abundant phospholipids such as PC or sphingomyelin processed by the other types of phospholipase C that these bacteria secrete. In B. cereus, there is experimental evidence that sphingomyelinase and PC-PLC, but not PI-PLC, are involved in phosphate retrieval [17, 18]. All the bacteria that produce PI-PLC are opportunistic pathogens, and there has been some interest in the potential role of the PI-PLCs as virulence factors [19–23]. Indeed, PI-PLC activity was first observed in studies of anthrax; the PIPLC secreted by Bacillus anthracis caused a large increase in plasma alkaline phosphatase early in infection [24]. Although we now know that this observation probably had little to do with the pathogenesis of anthrax, it did lead to the discovery of PI-PLC in other Bacillus spp. More recent studies with S. aureus, L. monocytogenes, and B. thuringiensis indicate that PI-PLC is often coregulated with other recognized virulence genes. PI-PLCs do not readily lyse cells because the mass abundance of their substrates in the cell surface is relatively low. PI is mainly located on the cytoplasmic surfaces of membranes and in most cells GPI is only a minor lipid. Furthermore, as noted above, these bacteria also produce at least one other type of PLC, which can hydrolyze relatively abundant cell surface phospholipids and as a consequence produce cell lysis more readily. While it seems likely that the function of PI-PLC is to degrade PI, free GPIs, and GPI-anchored proteins on the extracytoplasmic surface of host cells, there is relatively little information at the molecular level on the relationship of this activity to pathogenesis.

GPI-Specific Phospholipase C A key factor in the discovery of the GPI anchor was the observation that the variant surface glycoprotein (VSG) of the protozoan parasite, Trypanosoma brucei, was rapidly converted from a membrane bound form to a soluble form following osmotic lysis [25, 26]. Conversion was subsequently shown to be the result of an endogenous GPI-specific phospholipase C that cleaved the diacylglycerol moiety from the GPI anchor [27–29]. Following cleavage by GPI-PLC, VSG can be recognized by CRD antiserum, which suggests a reaction mechanism closer to that of bacterial PI-PLC than to mammalian PI-PLC (see earlier section, and [12]). VSG is extremely abundant for a cell surface protein (앒107 molecules/ cell) and was the source of the first GPI anchor to be thoroughly characterized. VSG has also proven useful because it is relatively easy to prepare substrates for GPI-PLC and GPI-PLD by biosynthetic labeling of the GPI anchor with [3H] myristic acid. The GPI-PLC from T. brucei has been purified and characterized


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extensively at the molecular level [30–35]. A recent revaluation of the substrate specificity of GPI-PLC has demonstrated that it has significant but relatively weak activity against PI [36], which has raised questions as to the natural substrate for GPI-PLC in the trypanosome. The picture is further complicated by the fact that the GPI-PLC is located on the cytoplasmic face of intracellular vesicles and consequently should hydrolyze PI or GPI precursors, but not GPI-anchored proteins such as VSG [29, 37]. The GPI-PLC has no obvious hydrophobic sequence or motifs for lipid anchor attachment, and the basis for its membrane association remains obscure. The physiological role of VSGs is to provide a protective surface coat against host defenses by means of antigenic variation. The ability of the trypanosome to switch expression among a repertoire of 앑1000 VSG genes (which have variable primary sequences but conserved secondary and tertiary structures) assures that there are always sufficient trypanosomes in the blood able to evade the immune response and maintain infection [38, 39]. Presumably, the role of GPI-PLC is to release VSG molecules at a reasonably rapid but in a controlled fashion in order to accommodate developmental changes during the life cycle, changing to a different coat protein in the insect host and back to VSG in the mammalian host and providing a mechanism to generate antigenic variation. However, the precise role of the GPI-PLC remains unclear [29]. The release of VSG during in vitro differentiation to the procyclic insect form involves proteolytic cleavage rather than GPI-PLC [40]. Furthermore, GPI-PLC null mutants are able to pass through a complete life cycle, indicating that GPI-PLC is not essential [41]. That mutants are also able to maintain persistent infection in mice indicates that the GPI-PLC is also not required for antigenic variation. However, the first peak of mutant parasitemia was reduced and the mice survived longer than in wild-type infections. One plausible explanation for these results is that the role of the GPI-PLC is to release the VSG rapidly when a trypanosome is under immune attack [29, 41]. The simultaneous release of massive amounts of VSG may provide a sort of ‘‘smoke screen’’ to allow trypanosomes expressing the same antigen to escape immune surveillance, along with those parasites which have switched expression to a different VSG gene. In support of this model is the observation that, in vitro, a variety of stress conditions will induce coat release without cell lysis [41–43]. Release of the 1,2-dimyristoylglycerol and inositol phosphoglycans, which are attached to the VSG, may also contribute to this diversionary tactic [44]. A gene encoding a GPI-PLC in T. cruzi has recently been cloned and the expressed enzyme has activity on GPI substrates [45]. The T. brucei and T. cruzi GPI-PLCs have 46% amino acid sequence identity with conservation of most residues predicted to be involved in substrate binding and catalysis. Comparative studies of these enzymes should be very informative because, unlike T. brucei, T. cruzi is an intracellular parasite in mammals. In contrast to substantial progress made with the trypanosomal GPI-PLC, attempts to identify a mammalian counterpart have proven less successful, and it is uncertain whether such an enzyme exists in higher eukaryotes. A preliminary report of sequence similarity between a bovine brain protein and the trypanosomal GPI-PLC has not been substantiated [46]. Apparent GPI-PLC activities were also reported in crude membrane fractions from liver and brain [47, 48].

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However, it is very likely that these activities were due to serum GPI-PLD, a major contaminant in homogenates prepared from tissues that were not perfused (see later section). As noted in other studies, phosphatidic acid released by the GPI-PLD can be rapidly converted to 1,2-diacylglycerol by PA phosphatase in cell lysates, giving the erroneous impression that initial cleavage was the result of a phospholipase C [49]. An apparent GPI-PLC activity was also purified from rat liver plasma membranes, but there are no recent reports of detailed molecular characterization of this enzyme [50]. GPI-PLD activity has been detected in liver plasma membranes and, for the reasons described above, may have been responsible for the initial degradation reaction [51]. Mammalian cells also contain large amounts of Ca2⫹-dependent phospholipase C activity which degrades the phosphoinositides (phosphatidylinositol and phosphatidylinositol phosphates); this activity is unlikely to interfere with GPI-PLC assays because none of this family of enzymes have been reported to act on GPI molecules (Table 11-1). A major impetus for attempts to isolate and characterize a mammalian GPIPLC was the proposal that such an enzyme would release inositol phosphoglycans from free GPIs or GPI-anchored proteins, as, for example, in response to insulin and a wide variety of other stimuli (for reviews of the extensive work in this area see [1, 52, 53]). Biological activities of the inositol phosphoglycans range from acute regulation of metabolism to cell growth and tissue differentiation. Inositol phosphoglycans with broadly similar chemical properties and biological activities can be produced in vitro by treating lipid fractions or GPI-anchored proteins isolated from a variety of cells with bacterial PI-PLC. However, attempts to purify these compounds have not yielded consistent biological activity profiles, chemical compositions, or physical structures. Consequently, after a decade of research there are still no strong leads for the synthesis of analogs which possess equivalent biological activities. An additional complication is the relatively recent observation that inositol phosphoglycans prepared with PI-PLC and GPI-PLD have similar potencies [44, 54]. Insulin may also stimulate release of GPIanchored proteins from cell surfaces by activation of a GPI-PLC [55–59]. Unfortunately, even though the same or a closely related GPI-specific phospholipase may be involved in both the generation of inositol phosphoglycans and protein release, these interesting findings have not led to its isolation or characterization.

GPI-Specific Phospholipase D Similar to the discovery of the trypanosomal GPI-PLC, a mammalian GPIdegrading activity was detected—before the formal description of its substrate, the GPI anchor! An endogenous enzyme activity was observed to remove the membrane anchor from alkaline phosphatase and convert the protein to a form no longer able to bind to membranes [60]. This activity required butanol to delipidate and solubilize the alkaline phosphatase (see later section). Ironically, this extraction procedure was used for many years (to prepare alkaline phosphatase in a ‘‘soluble’’ form that could be analyzed by nondenaturing gel electrophoresis) in complete ignorance of its mechanism. Initially the anchor-degrading activity was believed to be due to a Ca2⫹-dependent phosphoinositide-specific phospholipase C (see Table 11-1), which was known to be abundant in mammalian tissues [60]. The observation that the activity was present in both the particu-


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late and supernatant fractions of cells, was inhibited by EDTA, and required an acidic pH for optimal activity perpetuated this belief [61–63]. However, the GPIdegrading activity had relatively high thermal stability compared to phospholipase C, and the product of GPI degradation was phosphatidic acid instead of diacylglycerol [64]. These two distinct enzymatic activities could also be readily defined by the use of the transition metal chelator 1,10-phenanthroline, which is unable to chelate Ca2⫹ and does not inhibit the phospholipase C, [65, 66]. Reexamination of its subcellular distribution indicated that the activity detected in the supernatant fraction of organs such as liver, heart, and brain was due to contamination by serum/plasma which contained high levels (compared to cells) of GPI-PLD [65–67]. The recognition that GPI-PLD was present in mammalian plasma facilitated its purification and its molecular characterization. The remainder of this chapter will describe our current knowledge of GPI-PLD

Biochemical Properties of Plasma GPI-PLD The availability of a relatively abundant source in the serum/plasma allowed the purification, molecular cloning, and biochemical characterization of GPI-PLD. However, it should be emphasized that there is no comparable information for intracellular GPI-PLD, an important issue because the plasma form of the enzyme has little or no activity under physiological conditions. This section focuses on the properties of the plasma enzyme; the relationship between plasma GPI-PLD and intracellular GPI-PLD is discussed in detail later.

Metal Ion Dependence The first biochemical characteristic of the mammalian anchor-degrading activity to be discovered was its sensitivity to inhibition by the divalent cation chelator EDTA [60]; this inhibitory effect was also produced by EGTA and could be blocked by the addition of excess Ca2⫹ ions, demonstrating that inhibition was due to chelation of divalent cations [64, 68]. However, it was subsequently shown that the transition metal chelator, 1,10-phenanthroline, which does not chelate Ca2⫹ or Mg2⫹, was a much more effective inhibitor of GPI-PLD than either EDTA or EGTA [65, 66]. These observations suggested that inhibition was due to removal of a transition metal and that the blocking effect of excess Ca2⫹ was a result of competition for limiting amounts of chelator. This possibility was confirmed when it was demonstrated that the inhibitory effect of EGTA could be blocked or reversed with substoichiometric amounts of Zn2⫹ [69]. The lack of any requirement for Ca2⫹ was finally established when it was shown that the endogenous metal ion–EGTA chelates could be removed from GPI-PLD by gel filtration and full activity restored with Zn2⫹-EGTA chelates [70]. Analysis of purified GPI-PLD for metal revealed the presence of approximately 10 bound zinc ions enzyme molecule [69]. Although this result is consistent with the inhibitory effects of specific chelators, the number of zinc ions is unusually high; most zinc-dependent metalloenzymes typically have only 1, 2, or 3 bound zinc ions [71]. Furthermore, it is not clear where such a large number of zinc ions would bind. Those proteins which can bind ⬎3 zinc ions usually contain

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multiple repeats of a stereotypical zinc-binding domain. Motifs like the zinc finger domain are easily recognizable because they contain many cysteine/histidine residues with characteristic spacing located within a relatively short contiguous stretch of polypeptide [72]. However, there are no such zinc-binding motifs in the GPI-PLD cDNA sequence, and while at least two cysteine residues are required to coordinate a single zinc ion, there are only 10 cysteine residues in the entire GPI-PLD molecule. The conclusion that Ca2⫹ ions were not required for enzyme activity was somewhat surprising because information derived from the polypeptide and cDNA sequence predicted the presence of four Ca2⫹ binding sites [73, 74], (similar sites are also found in integrin-움 subunits [75]). The predicted binding sites in both GPI-PLD and the integrins show a strong relationship to the EF-hand motif found in numerous Ca2⫹-binding proteins, such as calmodulin, parvalbumin, and Troponin C (Fig. 11-2; [72]). However, the ‘‘integrin EF-hand’’ sites in both GPI-PLD and integrins have two additional features which distinguish them from the classic EF-hand motif [72, 75]: (i) they lack a glutamate residue at position 12 and (ii) there is a short, highly conserved and relatively hydrophobic stretch following the last coordinating residue (positions 10–15). The integrin EF-

Figure 11-2 GPI-PLD contains integrin EF-hand Ca2⫹ binding sites. The consensus sequence for the EF-hand motif found in many calcium-binding proteins is shown on the left. Conserved residues which are involved in coordination of the Ca2⫹ion are enclosed in shaded boxes. The residue at position 7 is not conserved because coordination is provided by the main chain carbonyl group; coordination at position 9 is via an H-bonded water molecule. Integrin EF-hand Ca2⫹-binding sites located in GPIPLD (repeats 1, 2, 3, and 6 and the integrin 움IIb subunit (repeats 5, 6, and 7) are shown on the right. Residues 1, 3, 5, 7, and 9 within these repeats are predicted to participate in Ca2⫹ coordination by comparison with the EF-hand consensus sequence. The bidentate coordination provided by the glutamate residue at position 12 in EF-hands is absent in GPI-PLD and the integrins. This residue is replaced instead by a short, highly conserved, and relatively hydrophobic segment (residues 10–15). The location of the ‘‘missing’’ coordinating residue in integrin EF-hands is not known, but may be contributed by relatively distant parts of the polypeptide.


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hand was originally predicted to directly participate in the adhesive interaction between integrin 움 subunits and their natural protein ligands (collagen, fibronectin and others); an acidic residue in the ligand was proposed to substitute for the missing coordination site at position 12. However, current models of integrin structure (see later section) do not implicate bound Ca2⫹ ions in direct interactions with ligand, and the source (if any) of the missing coordination sites is unknown. At present it seems most likely that the bound Ca2⫹ ions have a structural rather than a regulatory role. In spite of the observation that Ca2⫹ ions are not required for activity, there is some evidence suggesting that the integrin EF-hand sites in GPI-PLD are indeed functional. GPI-PLD has been shown to bind approximately five Ca2⫹ ions with a Kd of approx 10⫺5 M. [69]. Binding appears to be relatively selective and cannot be blocked by several other divalent cations, including Mg2⫹, Zn2⫹, Co2⫹, Mn2⫹, and Ni2⫹. Ca2⫹ ions may also play an important role in the maturation of newly synthesized GPI-PLD [70], as mutation of either residue 1 or 3 in any of the predicted Ca2⫹-binding sites resulted in a ⬎90% decrease in the amount of GPI-PLD secreted by transfected COS cells, and for one of these sites, GPIPLD secretion was completely abolished by the mutations. However, for the other sites, residual GPI-PLD had a similar specific activity to the wild type. This indicates that the mutations resulted in impaired secretion of GPI-PLD with normal activity rather than secretion of normal amounts of inactive GPIPLD [70]. The precise reason for impaired secretion is unknown and could result from several different causes: Ca2⫹-binding sites may be required for the initial folding of GPI-PLD, for the formation of oligomers, or for transit through the endoplasmic reticulum/Golgi.

Oligomerization When unfractionated serum or plasma are analyzed by gel filtration chromatography, GPI-PLD behaves as a particle of molecular mass approximately 500 kDa, which was initially attributed to the ability of purified GPI-PLD to aggregate or to associate with lipoproteins, particularly in the absence of detergent [65]. Subsequently, most GPI-PLD activity in human and bovine serum/plasma was shown to be associated with the HDL fraction [76, 77]. The exact physical nature of this interaction is unknown but appears relatively specific, as immunoaffinity chromatography binding of GPI-PLD is restricted to the LpA-I subfraction of HDL [77]. A minority of high-density lipoprotein (HDL) particles have been reported to contain protein molecules in addition to apoA-I and apoA-II, and the interaction between HDL and GPI-PLD could involve one of these other HDL-associated proteins. Alternatively, purified, delipidated apoA-I has been shown to disaggregate GPI-PLD, suggesting a direct interaction [78]. However, a large molar excess of apoA-I was required to disaggregate GPI-PLD, possibly due to the absence of the lipids which provide the normal milieu for apoA-I. Further studies using reconstituted HDL particles are required to define the specificity, and thus the potential physiological significance, of the GPI-PLD/ apoA-I relationship (see later section). A molecular mass of 100–120 kDa has been determined for GPI-PLD (purified from bovine or human plasma) by SDS-PAGE, compared to a value of 90.2

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kDa predicted from the bovine liver cDNA sequences [74, 76, 79]. The differences in these values presumably reflect glycosylation of the secreted protein, inferred from concanavilin A (Con A) and wheat germ lectin-binding studies and from sensitivity to N-glycosidase treatment [74, 76, 80]. However, as noted in the preceding paragraph, gel filtration studies indicate that purified GPI-PLD from bovine and human serum aggregates. The protein is also reported to be amphipathic and to bind relatively strongly to hydrophobic chromatography matrices such as phenyl and octyl sepharose [76, 79]. In the presence of Triton X-100, the molecular mass is approximately 200–350 kDa, which increases to ⬎400 kDa in the absence of detergent [69, 74, 76]. An exact molecular interpretation of these values is not possible given the inaccuracies inherent in size estimation by gel filtration and SDS-PAGE, but one possibility is that the GPI-PLD is a stable dimer which aggregates in the absence of detergent. Alternatively, the dissociated form of GPI-PLD might be an asymmetric monomer rather than a dimer, which would account for an anomalously high estimate of molecular mass by gel filtration [76]. GPI-PLD isolated from thrombin-treated human plasma is monomeric, even in the absence of detergent [79]. This interesting result has not been confirmed, and the appearance of a monomeric form of GPI-PLD may merely be a relatively trivial consequence of differences in isolation and analytical procedures.

Structure of GPI-PLD—Experimental Studies The limited structural information available for GPI-PLD derives from tryptic fragmentation studies [69, 80]. Controlled trypsin treatment cleaves GPI-PLD into three fragments (Fig. 11-3): two 앑40-kDa fragments, (A and B) and a 앑30-

Figure 11-3 Cleavage of GPI-PLD by trypsin. Native GPI-PLD (top of figure) is cleaved sequentially into three fragments by trypsin (arrows). Cleavage sites were located by N-terminal sequencing of the peptide fragments. Cterminal residues (italic numbers) were deduced from the location of predicted tryptic cleavage sites and from the size of the peptide. Also indicated are the locations of the epitope for mAb 191 (heavy shading), predicted Nlinked glycosylation sites (solid circles), and the 웁-propeller domain (light shading, see Fig. 11-5).


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kDa fragment (C). Of particular significance was the observation that tryptic fragmentation did not inactivate GPI-PLD; to the contrary, depending on the assay conditions, the catalytic activity of the trypsinized enzyme was either unchanged or increased from three- to 10-fold [69, 80], nor did trypsin treatment alter the inability of GPI-PLD to act on membrane substrates (see later section). The sensitivity of catalytic activity to inhibitors such as Lipid A, phosphatidic acid, EDTA, EGTA, or 1,10-phenanthroline or to thermal denaturation was also unaffected by trypsin treatment [69]. A comparison of N-terminal sequence analyses, the molecular mass of the tryptic fragments, and the effect of N-glycosidase treatment with the location of potential N-glycosylation or tryptic-cleavage sites indicated that fragments A, B, and C encompass residues 1–275, 326–589, and 590–816 (the C terminus). Kinetic studies with different incubation times or concentrations of trypsin showed that cleavage occurred in three stages (Fig. 11-3; [69, 80]): (i) cleavage at residue 590 occurs first to produce a 앑30-kDa C-terminal fragment (C); (ii) the remaining N-terminal fragment is then cleaved at several sites between residues 296 and 326 (i.e., at residues 296, 303, and 326) to produce a 앑40-kDa fragment (B) and a 앑50-kDa fragment (X); (iii) finally, the C terminus of fragment X is further degraded, probably at residue 276, to produce a 앑40-kDa fragment (A). The final reaction is detectable because of the relatively large decrease in molecular mass (probably as a result of glycosylation at residue 284), and this region also contains the epitope for mAb 191 (Fig. 11-3). The conversion of fragment X to fragment A is accompanied by complete loss of immunoreactivity. Whether additional cleavage occurs at the C termini of fragments B and C is unknown. The A, B, and C fragments of trypsinized GPI-PLD cannot be separated without prior denaturation [69, 80]. However, in the presence of 6 M urea, fragments A, B, and C can be separated by chromatography and renatured [80]. Although recovery following renaturation was low, activity could be detected in fragment A, whereas fragments B and C neither contained activity themselves nor enhanced the activity of A. These results suggest that the active site of GPIPLD is located in the N-terminal one-third of the protein. The remaining twothirds of the molecule is not required for activity, although it may enhance catalytic activity if it contains part of the substrate binding site (see later section and [81]). Unfortunately, attempts to confirm these data by expression of truncated forms of GPI-PLD containing only the N-terminal region in COS cells have not been successful. Deletion mutants based on the tryptic cleavage sites are not secreted or retained in the cell, suggesting that the individual fragments are unable to fold efficiently [70].

Structure of GPI-PLD—Modeling Studies The presence of similar multiple Ca2⫹-binding sites in GPI-PLD and in integrin움 subunits suggested the possible presence of other structural similarities between these two very different types of protein. Modeling studies indicated that the Cterminal region of GPI-PLD possesses seven FG . . . GAP repeats, each containing 앒60 resides, as already established for the N-terminal region of integrin 움 subunits [82]. GPI-PLD appears to be the only protein with such a strong resemblance to this region of the integrin-움 subunit. A major difference was the

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location of the predicted Ca2⫹-binding sites: in GPI-PLD these occur in repeats 1, 2, 3, and 6, whereas in integrins they usually occur in repeats 5, 6, 7, and occasionally in repeat 4 (Fig. 11-4). More recently, there has been an attempt to predict the folding pattern of the seven repeats in integrins and GPI-PLD [83]. In the absence of sufficient intrarepeat disulfide linkages to stabilize them, the repeats are too small to fold as independent domains. Consequently, the repeats were proposed to form a single, seven-bladed, 웁-propeller domain with the Ca2⫹-binding sites located in loops projecting from the base (Fig. 11-5). Each blade of the propeller, consisting of a four-stranded antiparallel 웁 sheet, is arranged around a central axis and makes extensive contacts with its two neighbors. In GPI-PLD, this domain is predicted to contain the residues from 350 to 816 (the C terminus). The model is compatible with tryptic fragmentation studies, in which the domain containing the catalytic site was located between residues 1 and 275 (see previous section). The model also predicts that residue 590 is located in an exposed loop projecting from the top of the 웁-propeller domain, consistent with its high sensitivity to tryptic cleavage (see previous section). Deletion mutants of GPI-PLD containing only the 웁-propeller domain are not expressed, even when contransfected with the N-terminal domain, suggesting that interactions between the two domains are essential for the initial folding [70]. The 웁-propeller model may also explain the sensitivity of GPI-PLD activity to small deletions at the C terminus; removal of 2–5 residues from the C terminus reduces the activity to approximately 30% of wild type [84], and removal of 6 and 7 residues reduces the activity to approx. 2 and 0% respectively. These changes coincide with deletions from the seventh blade of the propeller (Fig. 11-5). The C and N termini of the 웁 propeller are adjacent in the seventh repeat, and deletion of C-terminal residues would likely perturb the interaction between the 웁 propeller and the N-terminal catalytic domain. In the latter studies, although the GPI-PLD was inactive, normal amounts of the polypeptide were secreted

Figure 11-4 Structural domains in GPI-PLD and integrin-움 subunits. The distribution of predicted structural domains is compared for GPI-PLD and a typical integrin-움 subunit. In GPI-PLD, the catalytic domain (no shading), containing the active site, is located in the N-terminal one-third of the protein. The rest of the polypeptide contains the sevenfold repeat structure (shaded areas). In integrin 움-subunits, the repeats are located at the N terminus and are connected to a C-terminal transmembrane domain by an intervening segment, which is relatively variable in length and sequence between different 움 subunits. In some integrins, an additional domain, the I domain, is inserted between the second and third repeats (not shown). Ca2⫹-binding sites are located in repeats 1, 2, 3, and 6 in GPI-PLD (solid shading). In integrin-움 subunits, these sites are located in repeats 5, 6, and 7, and occasionally in repeat 4.


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Figure 11-5 웁-propeller structure of GPI-PLD. (Top) The folding predicted by the 웁-propeller model is shown for GPI-PLD. Each of the repeats (W1–W7) consists of a twisted, four-stranded antiparallel 웁 sheet. The major tryptic cleavage sites (arrows) are outside the 웁-propeller domain or in a loop projecting from the upper surface of the 웁 propeller. (Bottom) A top view of the 웁 propeller to illustrate the arrangement of the repeats (or ‘‘blades’’) around a central cavity. The first strand in each blade is located toward the center and the fourth strand is on the periphery as indicated by the strand numbers given for W1. As a result, polypeptide loops connecting the repeats pass from the periphery of W1, over the top surface, to the central aspect of W2. The polypeptide is contiguous within all repeats except for W7, where strand 4 is contributed by the N-terminal region and strands 1–3 by the C terminus (not shown). The Ca2⫹-binding sites found in W1, W2, W3, and W6 are located at the bottom of the 웁 propeller in loops between the first and second strands (filled circles).

and gross misfolding and retention by the cell apparently were not responsible for reduced catalytic activity [84]. An interesting feature of the 웁-propeller model that may have great functional importance is the prediction of a cavity in the center of the upper surface [83]. The lower part of this cavity is relatively apolar and contains no charged residues and could be appropriate as the location of the substrate binding site. The hydrophobic acyl chains of the GPI molecule could be shielded from water in the bottom of the cavity, with the polar head group projecting from the top. Another structural feature which may participate in substrate binding is the presence of numerous basic residues located around the cavity’s rim, which could be involved in binding of the negatively charged phosphodiester group. The location of a substrate binding site in the upper surface is consistent with the observation that enzyme activity is not affected by Ca2⫹ ions [69, 70]: the Ca2⫹ binding sites are proposed to be on the opposite face of the 웁-propeller.

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Inhibitors As described above, GPI-PLD is sensitive to inhibition by divalent cation chelators, presumably as a result of their ability to remove the bound zinc ions that are required for activity (see earlier section). However there are several other, nonchelating compounds that can inhibit GPI-PLD. The ability of purified GPIPLD to act on its GPI substrate in liposomes was inhibited by inclusion of dimyristoyl phosphatidic acid (PA) into the liposomes [85], on effect also observed when the substrate and inhibitor were dissolved in a large excess of detergent. Inhibition was not strictly dependent on the number or length of the fatty acid chains, provided that the total number of methylene groups was 14 or greater [86]; both dioctanoyl PA and palmitoyl lysoPA (LPA) had IC50 values of approximately 1–2 애M. Many other glycerolipids, phospholipids, and watersoluble phosphomonoester compounds were not inhibitory, suggesting that inhibition required both the phosphomonoester group and acyl group(s). Initially it was thought that PA activity was an example of product inhibition, exaggerated by the relatively low substrate concentrations used in standard GPI-PLD assay procedures. Later, Lipid A (the lipid moiety used by gram-negative bacteria to anchor the surface lipospolysaccharide coat) was also shown to be as inhibitory as PA and LPA [86]. Lipid A, as a disaccharide consisting of two glucosamine phosphate residues which are each acylated at the 2- and 3-positions, contains two moieties which resemble PA in the relative placement of phosphate and acyl groups. Monosaccharide analogs of Lipid A, as well as analogs of PA in which the glycerol backbone or the acyl groups are modified, were also inhibitory [81]. The mechanism by which Lipid A, PA, and LPA block GPI-PLD is not known, but the hydrophobic central cavity in the 웁 propeller is surrounded by basic residues that may bind these anionic molecules and prevent substrate interaction (see later section). The only other compounds that have been shown to inhibit GPI-PLD at micromolar concentrations are two azo suramin analogs [87]. The most effective of these (4,4⬘-bis(8-hydroxy-5-sulfo-7-quinolineazo)diphenyl sulfone; compound III) has an IC50 of 3.7 애M, but the mechanism of inhibition was not investigated. There is no obvious similarity between compound III and Lipid A, PA, and LPA except that compound III is anionic and consequently could bind to the central cavity in the 웁-propeller on GPI-PLD, as described above. 8-Hydroxyquinoline-5-sulfonate is a zinc chelator, and possibly these two moieties in compound III are simply removing bound zinc ions from GPI-PLD [87]. Bicarbonatecontaining buffers have also been reported to inhibit GPI-PLD activity in serum and liver membranes; inhibition was reversible and thought to result from carbamoylation of an amino group on the GPI-PLD. Sodium cyanate inhibition of GPI-PLD is consistent with this mechanism [48].

Distribution of GPI-PLD in Tissues and Cells GPI-PLD has been detected in a variety of cells and tissues (Table 11-2), as inferred from immunohistochemistry, in situ hybridization, Northern blotting, and assays of enzyme activity. The antibodies, oligonucleotide probes, and func-

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TABLE 11-2 Localization of GPI-PLD in Tissues and Cells Tissue/cell type

Detecton technique a

Reference

Cells b Pancreatic islet cells Neurons Mast cells Keratinocytes Leukocytes Bone marrow cells

IMM; GPI; RNA IMM IMM, in situ IMM; GPI IMM; GPI; RNA GPI; RNA

[49, 90, 92] [118] [89, 97] [49] [91, 99] [119]

Tissues Lung c Liver c Adrenal c Brain Skin—dermis c Skin—epidermis d Forestomach/esophagus d Stomach Skeletal muscle Heart Testis Coronary artery e

IMM; in situ RNA; in situ In situ IMM; RNA IMM IMM IMM; GPI IMM RNA RNA RNA IMM

[89, 97] [96–98] [97] [96, 98, 118] [89] [49] [49] [89] [96] [96] [96] [99]

a

IMM, immunohistochemistry; GPI, GPI-degradation assay; RNA, Northern blotting/RT-PCR; in situ, in situ hybridization. b Includes observations made both on tissues and in cells in culture. c–e Exclusively localized in c mast cells; d keratinocytes; e macrophages of atherosclerotic coronary arteries.

tional assays used were designed to detect plasma (human or bovine) GPI-PLD or its mRNA, because this is the only form that has been purified. There are major differences in the cell and tissue distributions shown with these techniques, which may reflect limitations in methodology or the existence of multiple distinct forms of GPI-PLD. Cellular forms of GPI-PLD have not been purified, and consequently there is no comparative biochemical data for plasma and cellular GPI-PLD. GPI-PLD activity has also been detected in milk and cerebrospinal fluid, but the amounts are ⬍1% of that found in plasma and it has been impractical to purify them from these sources. There is also relatively little information on the species distribution of GPI-PLD. Nevertheless, similar activities have been found in the serum/plasma of all mammals studied, as well as in chicken plasma and neural tissue [65, 88].

Immunohistochemical Studies Monoclonal antibodies have been raised against purified GPI-PLD from human (for example, 612C) and bovine GPI-PLD (191) and used in surveys of human and rat tissues, respectively (Table 11-2), where staining was relatively sparse [49, 89]. Binding to mAb 612C was found in several human tissues (lung, stomach,

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and dermis) but always restricted to mast cells [89]. In rat, staining with mAb 191 was mainly localized to keratinocytes in the stratified squamous epithelium of the forestomach, esophagus, skin, and hard palate [49]. Staining for mAb 191 was also observed in scattered individual motor and sensory neurons in the brain, spinal cord, and dorsal root ganglia, in smooth muscle cells, and in the islets of Langerhans. In contrast to humans, there was no evidence of mast cell staining in the rat gastrointestinal tract, lung, or dermis. Apart from differences in fixation and staining techniques, there are two main reasons which could account for the different results obtained in these species. First, the two antibodies may recognize different epitopes: mast cells could express a distinct isoform of GPI-PLD which is recognized by 612C but not by 191, and the neurons and keratinocytes a form that binds to 191 but not to 612C. That two genes encode distinct GPI-PLD isoforms has recently been questioned (see later section). Second, from in vitro experiments, the epitope for mAb 191 is known to be sensitive to tryptic degradation of GPI-PLD under conditions which leave the majority of the protein intact and do not destroy its catalytic activity (earlier section and Fig. 11-3). Consequently the same GPI-PLD might be recognized by 191 in some cells but not in others as a result of cell-specific cleavage by an endogenous protease.

Distribution of GPI-PLD Enzyme Activity For most tissues, direct measurement of GPI-PLD activity in homogenates or crude subcellular fractions is uninformative [49]. Even after in vivo perfusion, contamination by GPI-PLD from serum is so pervasive as to mask differences among tissues. The one exception is the keratinocytes in the stratified squamous epithelium of the rat forestomach, where it was possible to correlate a region of intense and highly localized staining with GPI-PLD activity in homogenates [49]. Detergent lysates of about 40 different primary cells or transformed cell lines (cultured in the absence of serum) have been analyzed for the presence of enzyme activity capable of degrading the model GPI-anchored protein substrate [3H]myristate-labeled variant surface glycoprotein from T. brucei ([3H]VSG). However significant degradation was observed only for a few cell types; these cells included transformed cell lines with the characteristics of 움 and 웁 cells from the islets of Langerhans, as well as a variety of myeloid cells and keratinocyte cell lines [49, 90, 91]. Secretion of GPI-PLD from cells was also observed from myeloid and 웁 cell lines but not from primary macrophages [90, 91]. Subcellular fractionation or immunofluorescence staining showed GPI-PLD located in secretory granules, but only in 웁 cells is there convincing evidence for physiological regulation of secretion [91, 92]. A major technical problem is the necessity for serum-free culture to prevent contamination with exogenous GPI-PLD [93], as specific activity and/or ‘‘concentration’’ of cell-associated GPI-PLD is generally comparable or lower than that observed in serum. Contamination is always a possibility. However, without careful characterization of each cell line, culture in serum-free media has the potential to induce changes in cellular metabolism, including unpredictable effects on GPI-PLD expression. Another difficulty is confirming that GPI-degrading activity is in fact due to GPI-PLD. As noted previously, [3H] myristate-labeled PA, which is produced by GPI-PLD, is immediately degraded to 1,2-diacylglycerol by extremely active intracellular PA phos-


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phatases, which gain access to the products of GPI degradation in the detergent lysate. Depending on the relative abundance of the GPI-PLD and PA phosphatase in a particular cell types, the [3H]PA may accumulate to detectable amounts [49, 90, 91]. This secondary reaction can erroneously register as GPI substrate degradation by a GPI-PLC. In keratinocytes, the situation is further complicated due to a phospholipase A2 which converts PA to LPA [49, 94], (avoidable by use of PA phosphatase inhibitors such as NaF, sphingosine, or sodium vanadate). Another important control is to inhibit GPI-PLD with 1,10-phenanthroline, to demonstrate that the first reaction was due to GPI-PLD and also to reveal secondary degradation [49, 91].

Distribution of GPI-PLD mRNA The cDNAs isolated from mouse pancreatic islet cells, bovine liver, human pancreas, and human liver have 74–95% sequence identity [73, 95, 96]. However, there remains some uncertainty about the number of genes. The cDNA sequences from human liver and pancreas predict numerous amino acid differences scattered fairly uniformly along the polypeptide, and humans may have two genes encoding GPI-PLD [95]. The single GPI-PLD gene in mouse (Gpldl) maps to the proximal region of chromosome 13 [96]. Sequence differences between liver and pancreas cDNAs could have resulted from a high degree of polymorphism or a relatively recent gene duplication in humans. Many of the amino acid differences may be due to sequencing errors because they derive from single base changes [96] Variability in sequences could complicate interpretation of mRNA distribution studies by in situ hybridization, Northern blotting, or RT-PCR, and tissue localization studies of GPI-PLD mRNA are confusing. In situ hybridization studies of bovine tissues were performed using a 400-bp cRNA probe derived from bovine liver cDNA [97]. Only in three of the 12 tissues examined was hybridization detected: adrenal gland, lung, and liver, and, again, localized exclusively to the mast cells. Although this work supported previous immunohistochemical studies implicating mast cells as a source of GPI-PLD, failure to obtain hybridization in other cell types was surprising. As described above, GPI-PLD had previously been detected in diverse cells, including pancreatic islet cells, bone marrow, keratinocytes, neurons, and leukocytes (see previous sections and, Table 11-2). The discrepancy between these results could have several explanations: (i) GPI-PLD secreted by the mast cells into the blood is taken up by other cells; (ii) mRNA levels are too low to be detected; or (iii) other cells express a distinct form of GPI-PLD that is not recognized by the cRNA probe [97]. In a recent Northern blot analysis of mouse tissues, using a full-length 움 cell cDNA as probe, a much broader tissue distribution was observed: Liver contained the most GPI-PLD mRNA but a robust signal was also obtained with brain, kidney, and skeletal muscle and sequence was detectable in heart, lung, and testis [96]. A characteristic pattern of transcript sizes (앑8.0, 5.4, and 3.9 kb) in different tissues was reproducible for liver and brain between individual mice. By contrast, in a separate study with human tissues, mRNA levels were comparable in brain and liver but were not detectable in kidney [97], and in human

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liver a 6.7-kb mRNA predominated whereas in bovine liver the only transcript observed was 3.9 kb [97, 98]. At present, it is unclear if the contradictory results of these three studies indicate genuine species differences or are methodological in origin. GPI-PLD mRNA has also been detected by Northern blot analysis of the J774 mouse-monocyte/macrophage cell line and by RT-PCR of primary human macrophages in confirmation of previous biochemical studies [91, 99].

Relationship between Plasma and Cellular Forms of GPI-PLD Until recently, a simple interpretation of the cell/tissue localization data would be GPI-PLD synthesis and secretion exclusively by mast cells, transported in the bloodstream/extracellular fluid, and uptake by other cells. The recent results of Northern blotting analyses, however, argue that several different cell types in multiple tissues can synthesize GPI-PLD. The technical problems described in previous sections severely reduce the sensitivity and reliability of methods for localizing GPI-PLD polypeptide and its mRNA, and negative results should be interpreted carefully. Nevertheless, liver as the principal source of GPI-PLD is consistent with clinical studies which indicate that the serum concentration of GPI-PLD is decreased in those liver diseases which affect hepatic synthetic ability (see later section and [100]). Whether particular cells synthesize their own GPI-PLD or depend on exogenous GPI-PLD also impacts on the potential subcellular site of GPI degradation as well as on the type of substrate. For example, endogenous GPI-PLD would have access to most components of the secretory and endocytic pathways, and its potential substrates would include not only cell surface proteins but also GPI molecules in the endoplasmic reticulum as diverse as newly synthesized GPIanchored proteins and free GPIs destined for attachment to proteins. By contrast, GPI-PLD synthesized exogenously would have access to GPIs at the cell surface and, if internalized by endocytosis, in the lysosomes, recycling compartments, and the trans-Golgi as well. Exogenous GPI-PLD would not have access to GPIs in the endoplasmic reticulum. GPI-PLD purified from serum can be internalized by neuroblastoma cells and retained by the cell for relatively long periods without being degraded [93]. GPI-PLD activity has also been detected in liver membrane fractions, where it may represent GPI-PLD molecules bound to ‘‘receptors’’ and in the process of being internalized. Detailed subcellular fractionation studies indicate that GPI-PLD is relatively enriched in the lysosomal fraction [101]. Taken together, these data strongly suggest that exogenous GPI-PLD can be internalized and gain access to intracellular compartments in a catalytically active form. As described earlier, the majority of GPI-PLD in plasma is bound to the LpA-I subpopulation of HDL particles with unknown physiological significance. Modulation of HDL levels by high fat and high cholesterol feeding had no effect on GPI-PLD levels [96], and GPI-PLD levels varied threefold between mice strains which had similar HDL and apoA-I levels [96]. The absence of any correlation between GPI-PLD and HDL may simply reflect the ability of apoA-I HDL to bind the amphipathic GPI-PLD molecule, allowing it to remain in the circulation at relatively high concentrations without aggregating. However, HDL could also be involved in the binding of GPI-PLD to plasma membranes


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and facilitate uptake into cells via receptor-mediated mechanisms. Also unknown is the nature of the interaction between GPI-PLD and lipoprotein particles, and whether it involves a specific interaction with apoA-I itself or with some other component (lipid or protein) within the same particle [77, 78]. The relative amounts of lipid and protein in the GPI-PLD/apoA-I HDL particles are also unclear because conflicting results have been obtained for their density [76, 77]. Other proteins are known to be associated with a minority of HDL particles—the identification of such proteins might be quite informative. Unfortunately, detailed molecular characterization of the GPI-PLD/apoA-I HDL particles is difficult because they represent a small minority (ⱕ0.2%) of total apoA-I HDL [77].

Physiological Role of GPI-PLD Despite the large amount of information regarding the biochemical properties of GPI-PLD, relatively little is known about its regulation or function. Obviously, GPI-PLD-mediated degradation of the GPI anchor is likely to release the attached proteins from the membrane, but for most proteins the purpose subserved by this process is not clear. As with many other areas of GPI research, the special role proposed for GPI-anchoring is uncharacterized and/or controversial, and not surprisingly, the consequences of anchor removal by GPI-PLD are even more obscure.

How Is GPI-PLD Regulated in Vivo? If GPI-PLD in plasma were to hydrolyze GPIs at maximal rate the enzyme would release most of the GPI-anchored proteins from the surfaces of blood and endothelial cells within a few minutes, a process that would be highly deleterious–cells stripped of surface GPI-anchored complement-regulatory molecules would be highly susceptible to bystander lysis similar to PNH red blood cells. Such a scenario does not occur because GPI substrates on intact cell surfaces or membranes are extraordinarily resistant to GPI-PLD, as compared to detergent-solubilized cells or purified GPI-anchored proteins used in in vitro assays [85, 102]. Lack of susceptibility does not appear to be the result of inhibitory molecules in the plasma: GPI-anchored proteins on intact cells or membranes are not only resistant to the action of GPI-PLD in plasma but also to the purified enzyme. Even though a large proportion of the GPI-PLD in plasma is bound to HDL, that interaction is probably not responsible for inactivity toward GPI substrates in cell surfaces (see earlier sections). When isolated GPI-anchored proteins are reincorporated into artificial membranes made from phosphatidylcholine, most (but not all) of the resistance to GPI-PLD exhibited by the native membrane can be regained [85, 102]. Fundamental structural features of the membrane make the GPI-anchored proteins embedded in it resistant to degradation by GPI-PLD. The ability of the membrane to influence enzyme activity is quite common among phospholipases and other lipid hydrolases [103, 104]. Indeed, for some enzymes, like the phosphoinositide-specific phospholipase C family (see Table 11-1), increased association with the membrane may play an important role in physiological regulation. The residual sensitivity exhibited by GPI-anchored

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proteins in artificial membranes could be due to their failure to faithfully mimic the surface of the plasma membrane [85]. The reported ability of GPI-anchored proteins at the cell surface to associate into sphingolipid/cholesterol enriched clusters or microdomains likely further reduces their susceptibility to GPI-PLD action [105] [see Chapter 10]. Indeed, disruption of these microdomains may be the reason why saponin (a cholesterol-binding agent) provokes GPI-PLD mediated release of Thy-1 from the surface of T cells [102]. Newly synthesized GPI-anchored proteins are not clustered into microdomains, presumably as a result of much lower concentrations of cholesterol and sphingolipids in the endoplasmic reticulum and cis-Golgi [106], and also explaining why cotransfection of GPI-PLD and GPI-anchored proteins into cells results in release of the GPI-anchored proteins [73, 107]. The site of degradation is believed to be intracellular because GPI-PLD that is secreted by the transfected cells is unable to release GPI-anchored proteins when added to the medium. Comparison of rates of degradation at intracellular sites with those obtained using in vitro assay systems is difficult, and it is not known whether the high rates observed in cells transfected with GPI-PLD is due to a relatively increased concentration of GPI-PLD in the lumen of the endoplasmic reticulum (which is dissipated upon secretion), the absence of microdomains in the endoplasmic reticulum membrane, or some more complex activation mechanism. An alternative has been suggested by the recent observation that alkaline phosphatase located in cell membranes was more susceptible to GPI-PLD isolated from baculovirus-infected insect cells compared to enzyme obtained from transfected CHO cells [98]. The biosynthesis of CHO cell and insect GPI-PLD indicated that the latter is secreted as an immature 98-kDa form in which terminal glycosylation did not take place; it is not known if the altered terminal glycosylation is responsible for the decreased activity toward cell membranes. However, the failure of a C-terminal antibody to bind to insect cell GPI-PLD, unless the antigen was first denatured, is consistent with glycosylation altering the conformation or accessibility of the C-terminal region. It will be important in the future to determine if the GPI-PLD isolated from baculovirus-infected insect cells also exhibits increased activity toward GPI-anchored substrates in artificial membranes. There is the intriguing possibility that, in mammals, GPI-PLD is only active at an intracellular location and is ‘‘inactivated’’ (by terminal glycosylation) during its passage to the cell surface. If so, then the relatively large amounts of GPI-PLD in plasma could be regarded as a residual protein in the process of being cleared from the circulation, and of little or no physiological significance.

Role of GPI-PLD in Release of GPI-Anchored Proteins from Cells and Tissues The spontaneous release of GPI-anchored proteins from tissues and cells has been widely observed—is GPI-PLD involved in the release process (Table 11-3). This simple question has proven very difficult to address experimentally due to fundamental technical problems. First, there are several alternative mechanisms which could ‘‘release’’ GPI-anchored proteins from cells, and to distinguish them, the cleavage sites must be identified by detailed analysis of the terminus of the glycan still attached to the released protein. Second, protein release is

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TABLE 11-3 GPI-PLD Mediated Release of GPI-Anchored Proteins from Cell and Tissues Source of released protein

Evidence in support of GPI-PLD mediated cleavage

ADP ribosyltransferase

Murine cytotoxic T cells

Alkaline phosphatase

Canine serum

Axonin-1

CD59 DAF

Chicken dorsal root ganglion cells Human urine HeLa cells

Heparan sulfate proteoglycan

Human bone marrow cells

NCAM

Murine myoblast cells Human cerebrospinal fluid

Release can be increased by physiological stimuli; 1,10-phenanthroline inhibits release Release may occur physiologically during enterohepatic circulation of bile acids; inositol analysis; no CRD reactivity; electrophoretic mobility [3H]Ethanolamine/[3H]inositol labeling; no CRD reactivity; 1,10-phenanthroline inhibits release Structure of C-terminal glycopeptide [3H]Ethanolamine labeling; no CRD reactivity; 1,10-phenanthroline inhibits release [3H]Ethanolamine labeling; no CRD reactivity; 1,10-phenanthroline inhibits release [3H]Inositol labeled product released by nitrous acid deamination Chromatographic behavior of C-terminal glycopeptide

Protein

Thy-1

Ref. [120]

[110, 112]

[88]

[121, 122] [123]

[119]

[124] [125]

generally slow and constitutive, so limited amounts of material are available for analysis, and secondary degradation reactions may obscure the original mechanism of release. Secondary degradation is a particular problem in the analyses of proteins isolated from body fluids, where the release rate is generally unknown and potentially highly variable. There are only two examples in which release of GPI-anchored proteins can be acutely increased by physiological stimuli (Table 11-3). Ironically, there is a relatively sensitive and specific technique for the identification of PLC-mediated cleavage of GPI-anchored proteins—but not for GPIPLD. The technique is based on the exposure of inositol 1,2-cyclic phosphate by both PI-PLC and GPI-PLC (Fig. 11-1). As described previously, this moiety is the dominant epitope for CRD antisera, which is raised by immunizing rabbits with PLC-cleaved GPI-anchored proteins (see earlier section). Proteins released from cells by an endogenous GPI-PLC can be detected by immunoblot with reasonable sensitivity and specificity; proteins released by GPI-PLD do not react with CRD antisera. There are several other release mechanisms, such as proteolytic cleavage, alternative splicing, vesiculation, and secondary degradation of PLC-released protein by a cyclic phosphodiesterase, which can all give rise to a negative CRD result. The first two of these can be studied by labeling with [3H]inositol, but [3H]inositol is a relatively poor label for GPI anchors because its incorporation into GPI precursors is slow. [3H]ethanolamine has been used as an alternative label because incorporation is faster and although it does not identify the cleavage site as precisely as [3H]inositol, distinction of phospholipase

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action from proteolysis is possible. Unfortunately, there are no specific inhibitors of GPI-PLD suitable for studying the release of GPI-anchored proteins. The PA and Lipid A analogs inhibitors (see earlier section) are relatively insoluble and difficult to introduce into intact cells. 1,10-Phenanthroline and its impermeant analog bathophenanthroline disulfonic acid, which have also been used to block release of GPI-anchored proteins, inhibit metalloproteases as well as GPI-PLD. These analytical limitations make it necessary to combine evidence from several techniques to provide convincing evidence for the involvement of GPI-PLD (see Table 11-3). Another potential complication is that GPI-anchored proteins are often enriched in small vesicles or other aggregates that are released from cells [108, 109]. Culture media must be ultracentrifuged or fractionated by gel filtration chromatography prior to analysis. GPI-anchored proteins released from cells in vesicles or aggregates may have increased susceptibility to extracellular forms of GPI-PLD. For example, the alkaline phosphatase in plasma is generally a mixture of hydrophobic oligomers, which still retain the GPI anchor, and hydrophilic dimers, which have been cleaved by GPI-PLD, and it has proven difficult to establish whether the hydrophilic dimers were produced by GPI-PLD before or after their release from the cell surface (see Table 11-3 and [110–112]).

What Is the Physiological Role of GPI-PLD Mediated Anchor Degradation? Although there is abundant evidence that GPI-PLD can degrade GPI-anchored proteins under physiological conditions, there is little information as to the function of GPI degradation. The following speculative discussion of potential roles is consistent with the known biochemical properties of GPI-PLD and GPI anchors, but there is little or no direct experimental evidence in support of any of them. Does GPI-PLD Generate Biologically Active Inositol Glycans? There are some data to indicate that the inositol glycan head group of GPIs has biological activity (see earlier section). Inositol glycans were initially proposed to result from the cleavage of free GPI molecules by a GPI-PLC in mammalian cells, on stimulation by insulin and other hormones and growth factors (for review see [52]). Despite considerable research, the structure of the inositol glycans and their subcellular sites remain uncertain, and there has been no progress in the molecular characterization of mammalian GPI-PLC. GPI-PLD might be involved in the origin of these molecules [54]. Does GPI-PLD Regulate Intracellular GPI Metabolism? GPI-PLD located in the endoplasmic reticulum could have an important role in degrading free GPI molecules there, in a process that might simply serve a ‘‘salvage’’ or ‘‘recycling’’ function for GPI precursors not used for attachment to proteins (or attached to misfolded proteins). Also possibile is that GPI-PLD could prevent free GPI molecules exiting the endoplasmic reticulum to gain unregulated access to the cell surface, especially important if cell surface GPI molecules participate in cellular signaling processes.


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Does GPI-PLD Regulate the Function of GPI-Anchored Proteins? Release of a GPI-anchored protein by GPI-PLD would be a very efficient and rapid mechanism for downregulating adhesion molecules or GPI-anchored proteins that act as ‘‘receptors’’ for extracellular ligands. As a consequence, GPI-PLD could rapidly disrupt cell adhesion or signaling mediated by these proteins. Does GPI-PLD Regulate Intercellular Transfer of GPI-Anchored Proteins? GPI-anchored proteins are known to transfer between cells in vivo, even if little is known about the rate and mechanism of this process. Rapid rates of transfer have been demonstrated in vitro between cells in contact [113]. GPIPLD could play a role in regulating or restricting this process in the bloodstream. GPI-anchored proteins transferred between cells in contact might escape degradation, whereas those engaged in fluid phase transfer would be susceptible to recognition and degradation by GPI-PLD. In essence, GPI-PLD could prevent random and promiscuous intercellular transfer of proteins between different cell types, except when sanctioned by close contact brought about by specific adhesion molecules.

GPI-PLD and Human Disease Even though the amount of GPI-PLD in humans is severalfold lower than in other mammals, there is still sufficient GPI-PLD in human serum to degrade approximately 20% of the GPI in the cell surface in 20 s. Massive release of proteins does not occur because the activity is restricted by mechanisms described in an earlier section. However, if constraints were to be disrupted, an obvious consequence would be the removal of complement regulatory molecules such as DAF and CD59 from the cell surface and the relatively rapid development of a ‘‘PNH-like’’ condition, which would affect not only most blood cells, but endothelial cells as well. More subtle effects on cell behavior could occur as a result of the removal of GPI-anchored molecules that are involved in cell signaling or adhesive interactions. Pathological increases in the rate of degradation of GPI-anchored proteins could occur by at least three distinct mechanisms: (i) even relatively subtle alterations of the membrane surface could cause loss of resistance to GPI-PLD (like disruption of microdomains); (ii) structural alteration of GPI-PLD (by defective glycosylation, for example) would overcome the resistance of the membrane [98]; (iii) increased secretion of GPI-PLD from cells due to abnormal regulation of GPI-PLD synthesis (as with oxidant stress; see below) could occur. Pathological suppression of GPI-PLD activity might also be deleterious. As described in an earlier section, GPI-PLD is inhibited by micromolar concentrations of LPA. Normally LPA concentrations in the plasma are very low, but following platelet activation, LPA concentrations in serum can be as high as 20 애M [114, 115], and be equally elevated in the plasma of patients with ovarian cancer [116]. Normal functions of GPI-PLD could be suppressed by LPA. Only two relatively limited studies compared the amount of GPI-PLD in the serum in patients and in healthy individuals [100, 117]. Activity was decreased in chronic liver diseases such as cirrhosis and hepatocellular carcinoma but increased in acute liver disease and pneumonia. GPI-PLD also decreased with age

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in normals. There was no correlation between GPI-PLD and alkaline phosphatase in the serum [100]. These results are broadly consistent with the properties of the target membrane as well as with the amount of GPI-PLD determining the rate of GPI degradation (see earlier section). A recent study implicated GPI-PLD in the inflammatory reaction that accompanies atherosclerosis [99]: immunohistochemistry of atherosclerotic coronary arteries revealed strong staining that was not present in normal arteries, and the staining was restricted to subsets of macrophages (including both foam and nonfoam cells) and did not occur in smooth muscle or endothelial cells. Of particular interest was the observation that GPI-PLD in atherosclerotic arteries was colocalized in macrophages that also expressed oxidation epitopes. Furthermore, in vitro studies showed that oxidant stress increased the amount of GPIPLD mRNA [99].

References 1. Low, M. G. (1989). The glycosyl-phosphatidylinositol anchor of membrane proteins. Biochim. Biophys. Acta 988, 427–454. 2. Low, M. G., and Brodbeck, U. (1999). Enzymes cleaving the phosphodiester bond in the GPI anchor. In ‘‘GPI-Anchored Biomolecules’’ (S. Ilangumaran and D. C. Hoessli, Eds.). R. G. Landes Company, Austin, TX. 3. Low, M. G. (1990). Degradation of glycosyl-phosphatidylinositol anchors by specific phospholipases. In ‘‘Cell and Molecular Biology of Membrane Proteins’’ (A. J. Turner, Ed.), pp. 35–63. Ellis Horwood, Chichester, England. 4. Low, M. G. (1992). Phospholipases that degrade the glycosyl-phosphatidylinositol anchor of membrane proteins. In ‘‘Lipid Modification of Proteins: A Practical Approach’’ (N. M. Hooper and A. J. Turner, Eds.), pp. 117–154. Oxford University Press, Oxford. 5. Gandhi, A. J., Perussia, B., and Goldfine, H. (1993). Listeria monocytogenes phosphatidylinositol (PI)-specific phospholipase C has low activity on glycosyl-PI-anchored proteins. J. Bacteriol. 175, 8014–8017. 6. Heinz, D. W., Ryan, M., Bullock, T. L., and Griffith, O. H. (1995). Crystal structure of the phosphatidylinositol-specific phospholipase C from Bacillus cereus in complex with myo-inositol. EMBO J. 14, 3855–3863. 7. Heinz, D. W., Ryan, M., Smith, M. P., Weaver L. H., Keana, J. F. W., and Griffith, O. H. (1996). Crystal structure of phosphatidylinositol-specific phospholipase C from Bacillus cereus in complex with glucosaminyl(움1씮6)-D-myo-inositol, an essential fragment of GPI anchors. Biochemistry 35, 9496–9504. 8. Moser, J., Gerstel, B., Meyer, J. E. W., Chakraborty, T., Wehland, J., and Heinz, D. W. (1997). Crystal structure of the phosphatidylinositol-specific phospholipase C from the human pathogen Listeria monocytogenes. J. Mol. Biol. 273, 269–282. 9. Heinz, D. W., Essen, L. O., and Williams, R. L. (1998). Structural and mechanistic comparison of prokaryotic and eukaryotic phosphoinositide-specific phospholipases C. J. Mol. Biol. 275, 635–650. 10. Iwasaki, Y., Tsubouchi, Y., Ichihashi, A., Nakano, H., Kobayashi, T., Ikezawa, H., and Yamane, T. (1998). Two distinct phosphatidylinositol-specific phospholipase Cs from Streptomyces antibioticus. Biochim. Biophys. Acta 1391, 52–66. 11. Volwerk, J. J., Shashidhar, M. S., Kuppe, A., and Griffith, O. H. (1990). Phosphatidylinositolspecific phospholipase C from Bacillus cereus combines intrinsic phosphotransferase and cyclic phosphodiesterase activities: A 31P NMR study. Biochemistry 29, 8056–8062. 12. Cross, G. A. M. (1979). Crossreacting determinants in the C-terminal region of trypanosome variant surface antigens. Nature 277, 310–312. 13. Zamze, S. E., Ferguson, M. A. J., Collins, R., Dwek, R. A., and Rademacher, T. W. (1988). Characterization of the cross-reacting determinant (CRD) of the glycosyl-phosphatidylinositol


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Appendix: Sequence of the Coding Region of the Human PIG-A Gene1

1

From Miyata et al. (1993). Science 259, 1318–1320. The GenBank accession number of PIG-A is D11466.

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Appendix: Sequence of the Coding Region of the Human PIG-A Gene

Index

A Acetylcholinesterase deficiency, in PNH erythrocytes, 88 N-Acetylglucosamine, in biosynthetic pathway of GPI, 201 Acidified serum lysis test, 11, 50–51, 64, 66, 127–128 ADP-ribosyltransferase, delivering T cell activation signals, 164–165 Animal models genetically engineered mouse, 119–120 human hematopoiesis in, 117–118 PNH chimeric mice bearing PIG-A(⫺) HSC, 149–154 mouse Piga gene, 141–143 PIG-A(⫺) ES cell differentiation in vitro, 143–149 Anisotropy, fluorescence, 228–230 Antibody CD52, 40–41 and complement, hemolytic action, 6 cross-linking, GPI-anchored proteins, 212–213 mediated clustering of GPI-anchored proteins, 159 monoclonal, see Monoclonal antibody monospecific, blocking MIRL function, 75 not found in PNH serum, 51 TSA-1, 162

Anticoagulation, in treatment of thrombosis in PNH, 109 Antigen density, and complement activation, 56–57 TSA-1, antibodies, 162 Antithymocyte globulin, toxicity in PNH patients, 129 Aplastic anemia autoimmunity in, 124–125 relationship to PNH, 14–15, 121–125 Apoptosis, resistance of HSC, 118–119 Asymmetry, transbilayer, membrane lipids, 222–223 Atherosclerosis, inflammatory reaction, GPI-PLD role, 262 ATIII deficiency, associated with venous thrombosis, 103 Autoactivation, C1r molecules, 57 B Basic fibroblast growth factor, glypican as binding site, 168 웁 propeller domain, GPI-PLD, 250–252 Biochemical properties, plasma GPIPLD, 245–252 Biological implications, GPI anchoring, 210–213 Biosynthetic pathway, GPI, 200–203

271

272

Index

Bone marrow chimeric mice with PIG-A(⫺) hematopoietic progenitors, 153–154 failure in PNH clinical course and treatment, 127–130 pathophysiology, 115–120 Bone marrow transplantation for PNH, 129–130 in treatment of PNH hemolysis, 83 Brain-derived immunoglobulin superfamily molecule-1, 165 Budd–Chiari syndrome, 12–13, 109, 130 C Calcium binding sites, GPI-PLD, 250–251 Calcium ions, GPI-PLD dependent on, 246–247 CAMPATH-1, see CD52 Caveolae enriched in molecules involved in signal transduction, 184–185 and GPI-anchored proteins, 169, 223–224 C1, binding to Fc portion of IgG or IgM, 55–57 C3b, factor B binding, 63 C4b-binding protein, 58 C5b67 complex, C8 binding, 60–61 C8-binding protein, blocking formation of MAC, 73–77 C9, polymerization, 61 C3 convertase activity of classical pathway, 58 DAF role, 80 regulation by erythrocyte membrane proteins, 69–73 C5 convertase alternative pathway, 63–64 classical pathway, 59–60 CD1 murine CD1d1, 165 restriction element, nonpolymorphic, 191 CD14, lipopolysaccharide receptor, 166–167 CD24, cross-linking, 163–164 CD48, monoclonal anti-sgp-60 antibody, 162–163 CD52 antibody, 40–41

cross-linking, 165 CD59, see also Membrane inhibitor of reactive lysis deficiency in PNH, 14 CD73, effect on interleukin-2 production, 163 Cell biology, mammalian, evidence for GPI-mediated signaling, 182–185 Cell lines, paired, in study of marrow failure in PNH, 118–119 Cells, see also Red cells; Stem cells apical surface and GPI-anchored proteins, 169–170 protein sorting to, 231–232 blood, genesis in embryo, 144–145 GPI-anchored protein release from, GPI-PLD role, 258–260 GPI-PLD distribution, 252–257 host regulation by ceramide-containing GPIs of T. cruzi, 189 signal transduction by protozoal GPIs, 185–187 PNH, artificial, 81–83 Cell signaling and GPI-linked proteins, 161–165 mediated by GPI-anchored proteins, 182–183 role of rafts, 232–233 Chimeric mice bone marrow, with PIG-A(⫺) hematopoietic progenitors, 153–154 with PIG-A(⫺) HSC, generation, 149–150 Cholesterol linking domains across lipid bilayer, 232–233 liposomes rich in, 227 Ciliary neurotrophic factor receptor, 183 Ciliary neurotrophic factor receptor 움, 165–166 Ciliated neurotrophic factor, 122 Clinical behavior, PNH, and pathophysiology of marrow failure, 115–119 Clinical course, marrow failure in PNH, 127–130 Clonal dominance, and Piga disruption, 150–151 Clonality, stem cell, 115

273

Index

Clustering, GPI-anchored proteins, 159, 224 Complement alternative pathway activation, 61–64 discovery, 53–55 alternative system of activation, 11–12 and antibody, hemolytic action, 6 classical pathway, 55–61 effect on platelet release of serotonin, 105–106 mediated lysis: PNH erythrocyte sensitivity to, 68–83 PNH erythrocyte sensitivity to, 13–14 red cell lysis affected by, 9 Complement lysis sensitivity assay, 64–65 Complement receptor type I activity on PNH erythrocytes, 70 recombinant soluble form, 84–85 short consensus repeats, 58 Complement-regulatory proteins, GPIanchored, 88 Compstatin, inhibitor of complement activation, 85 Corrals, constraining lipid-linked molecules in membrane, 228 Corticosteroids, effect on PNH hemolysis, 83–84 Cre/loxP system, Piga conditional disruption by means of, 151–153 Crosby, W. H., relationship of thrombosis and PNH, 12 Cross-reacting determinant, antiserum, 242 Culture systems PIG-A(⫺) embryoid bodies grown in, 147 In vitro colony culture studies of marrow failure in PNH, 116–117 Cyclosporin A in EBV transformations of PNH PBL, 126 in therapy for PNH, 129 Cytopathic stem cell-tropic virus, 123 D Dacie, John Vivian, description of hemolysis in acidified serum, 9–11 DAF, see Decay-accelerating factor Deacylation, inositol ring, in GPI biosynthetic pathway, 202

Decay-accelerating factor and artificial PNH cells, 82–83 deficiency in PNH, 70–72 isolated, 79–81 effect on GPI anisotropy, 230 identification, 13–14 protection of erythrocytes, 62 quantitated on PNH phenotypes, 78 restriction of convertase activity, 58 Decoy death receptor, 118 Degradation GPI-anchor, role of GPI-PLD, 260–261 GPI-anchored proteins by GPI-PLD, 257–258 Deletions, large, PIG-A, 33 Detergent, resistant membranes: connection with rafts, 223–227 Diacylglycerol, replacing alkylacylglycerol, 202 Diagnosis, PNH, 8–15 Dimyristoyl phosphatidic acid, inhibitor of GPI-PLD, 252 Disease, human, GPI-PLD and, 261–262 DNA, genomic vs. complementary, PIG-A mutations, 27 Dominance test, pattern of inheritance of PNH abnormality, 22–23 E EF-hand motif, GPI-PLD, 246–247 Embryo, blood cell genesis, 144–145 Embryoid bodies cell differentiation in, 145–146 formed by PIG-A(⫺) ES cells, 146–147 Embryonic stem cells Piga gene inactivation in, 142–143 PIG-A(⫺), in vitro differentiation, 143–149 Endoplasmic reticulum GPI biosynthetic pathway steps taking place on, 203 GPIs in, 256 Endosomes, GPI-anchored protein retention in, 231–232 Environmental factors, precipitating PNH attack, 4–5 Enzyme activity, GPI-PLD, distribution, 254–255 Erythrocyte membrane proteins inhibiting MAC, 73–77 regulation of C3 convertase, 69–73

274

Index

Erythrocytes lipid bilayer, disruption of integrity, 61 PNH, see PNH erythrocytes ES, see Embryonic stem cells Ethanolamine phosphate, modification of Man1, 201–202 Experimental studies, GPI-PLD structure, 248–249 F Factor B, activated by factor D, 62–63 Factor D, activation of factor B, 62–63 Fc moiety, IgG and IgM, C1 binding, 55–57 Fibrinolysis, cell-mediated, and uPAR, 108–109 Fluorescence anisotropy, 228–230 Fluorescence recovery after photobleaching, 227–228 Fluorescence resonance energy transfer, 228–229 Folate-binding protein, GPI-anchored, 167–168 Folate receptor, GPI-anchored or chimeric transmembrane-anchored, 229–230 Frameshift mutations, PIG-A, 33 FVIII, mutations, 42–43 G GAA1, homologs, cloning, 206–207 Genetics, biochemical, PNH in somatic cells, 21–23, 43–44 Glycoprotein, variant surface, and GPIspecific phospholipase C, 242–243 Glycosylphosphatidylinositol, see GPI Glypicans, GPI-anchored heparin sulfate proteoglycans, 168 gp130 HSC expressing, 122–123 uPAR association with, 167 G6PD deficiency, 42–43 GPI biosynthetic pathway, 200–203 ceramide-containing, of T. cruzi: host cell regulation by, 189 derived mediators of hormone action, 183–184 intermediates, chemically synthesized, 199–200 parasite, as functional toxins and pathogenicity factors, 187–188

protozoal, insulin-mimetic activities, 189–190 GPI-anchor addition to proteins, 207–210 cleavage by specific phospholipases, 239 defective mutants: mammalian and yeast, 203–207 degradation, role of GPI-PLD, 260–261 function, and raft hypothesis, 221–222 role in protein targeting to TIMs, 211–212 GPI-anchored proteins ⫹ and ⫺ progenitors, 117 antibody cross-linking, 212–213 association with caveolae, 169, 223–224 clustering, 159, 224 DAF, 59 detergent insolubility, physicochemical basis, 226–227 function as receptors, 166–168 regulatory role of GPI-PLD, 261 immunity to, and CD1d-restricted NK T cells, 190–191 lateral mobility, 168–169 higher in plasma membrane, 210–211 lipid-dependent retention in endosomes, 231–232 in neural tissues, 165–166 newly synthesized, 258 in parasites, 179–180 release from cells and tissues, GPIPLD role, 258–260 role in insulin action, 166 lymphocytes, 161–165 signaling mediated by, 182–183 GPI-anchored receptors, on HSC, 121–122 GPI-like structures, in parasites, 180–182, 191–192 GPI-PLD distribution in tissues and cells, 252–257 inhibitors, 252 metal ion dependence, 245–247 oligomerization, 247–248 physiological role, 257–262 structure experimental studies, 248–249 modeling studies, 249–251

275

Index

GPI-signal sequences, 207–210 Granulocytes PNH, mosaic, 67 survival, 15 Gull, William, first description of PNH, 2–3

H Ham, Thomas Hale, early experiments on hemolysis, 8–9, 11 Ham test, see Acidified serum lysis test Hematopoiesis human, in animal models, 117–118 impaired, amelioration, 129 PNH patients, 15 SCID mice, by cells from PNH patients, 154–155 Hematopoietic stem cells GPI-anchored receptors, 121–122 PIG-A(⫺), chimeric mice bearing, 149–154 PIG-A mutant, 44, 114–115 pluripotent, 87–88 therapeutic transplant, 115–116 Hemoglobinemia, relationship to sleep, 51–53 Hemoglobinuria march, 5, 8 PNH, as epiphenomenon, 49–50 relationship to sleep, 51–53 Hemolysis in acidified serum, description by Dacie, 9–11 early experiments by Ham, 8–9, 11 PNH, treatment, 83–86 Heteroduplex analysis, PIG-A mutations, 28 Heterogeneity, lateral, in cell membranes, 222–223 High-density lipoproteins, interaction with GPI-PLD, 247 Hijmans van den Bergh, A. A., description of PNH, 6–7 Homologous restriction factor, blocking formation of MAC, 73–77 Hormone action, GPI-derived mediators, 183–184 HSC, see Hematopoietic stem cells Hypoventilation, during sleep, and hemoglobinemia, 51–53

I Immune model, PNH pathogenesis, 120–127 Immunity, to GPI-anchored proteins, and CD1d-restricted NK T cells, 190–191 Immunoglobulins, IgG and IgM, C1 binding, 55–57 Immunohistochemical studies, GPI-PLD localization, 253–254 Inositol 1,2-cyclic phosphate, GPIanchored proteins cleaved by, 241–242 Inositol glycans, biologically active, GPIPLD role, 260 Inositol phosphoglycans, release from free GPI-anchored proteins, 244 Insulin action: role of GPI-anchored proteins, 166 mimetic activities of protozoal GPIs, 189–190 Interleukin-2, production, effect of CD73, 163 In vitro colony culture studies, marrow failure in PNH, 116–117 Iron, oral therapy for PNH hemolysis, 84 K Keratinocytes, GPI-PLD enzyme activity, 254–255 Knockout mouse PIG-A, 119–120 uPAR, 109 L Landois, Leonhard, transfusion experiments in animals, 5 Lateral mobility, GPI-anchored proteins advantages, 168–169 higher in plasma membrane, 210–211 role in positioning, 161 Leishmania glycoconjugates, regulation of macrophage function, 188–189 GPI-like structures, 181–182 Lipid A, inhibitor of GPI-PLD, 252 Lipid remodeling, in GPI biosynthetic pathway, 202–203 Lipophosphoglycan, Leishmania, 188–189 Lipopolysaccharide receptor, CD14, 166–167

276

Index

Liposomes, cholesterol and sphingolipidrich, 227 Liquid-ordered phase, promotion in artificial membranes, 226–227 Liquid-ordered regions, lipid bilayers, 223 Lymphoblastoid cell lines, PNH phenotype, 21 Lymphocytes GPI-anchored protein(⫺), 120–121 GPI-anchored protein role, 161–165 from PNH patients, exerting GPIdependent selection, 126–127 M MAC, see Membrane attack complex Macrophage function, regulation by glycoconjugates of Leishmania, 188–189 Mannosyl residues, in biosynthetic pathway of GPI, 201–202 Marchiafava, E., clinical description of PNH, 6–7 Membrane attack complex abnormal regulation on PNH erythrocytes, 72 formation, 60–61 inducing lysis of PNH III cells, 68 inhibition by erythrocyte membrane proteins, 73–77 Membrane inhibitor of reactive lysis C8 binding, 61 dysfunctional on artificial PNH cells, 82–83 isolated deficiency in PNH, 79–81 quantitated on PNH phenotypes, 78 recombinant soluble form, 85 and role of HRF/C8bp, 74–75 Membrane microdomains, GPIs, 184–185 Membranes cell, lateral heterogeneity in, 222–223 detergent-resistant: connection with rafts, 223–227 living cell, evidence for rafts, 227–230 Membrane structure, and GPI anchors, 168–170 Messenger RNA, GPI-PLD, distribution, 255–256 Metabolism, GPI, regulatory role of GPIPLD, 260 Metal ions, GPI-PLD dependent on, 245–247

Methylcellulose, media containing, PIGA(⫺) embryoid bodies grown in, 147 Microvesicles, platelet and red cell, role in PNH, 107 Mini transplant, therapy for PNH, 130 MIRL, see Membrane inhibitor of reactive lysis Modeling studies, GPI-PLD structure, 249–251 Molecular basis, erythrocyte phenotypes of PNH, 77–78 Monoclonal antibody humanized, 85 Ly-6E, 162 study of GPI-PLD localization, 253–254 Mutants gp1, gp2, and gp3, 205 GPI-anchor defective, 203–207 Thy-1 negative lymphoma, 205, 208 Mutations PIG-A, see PIG-A mutation X-linked genes, comparison, 42–43 N Natural history, PNH, 127–128 Neural tissues, GPI-linked proteins in, 165–166 Nonreceptor tyrosine kinase lipid-linked, activation, 232–233 and presence of rafts, 223–224 O Oligomerization, GPI-PLD, 247–248 웆 site, GPI-signal sequence, 207–208, 210 P Painting clinical potential, 222 reengineering of cell surfaces as, 213 Palmitoyl lysophosphatidic acid inhibitor of GPI-PLD, 252 serum levels in disease, 261 Parasites, GPIs, as functional toxins and pathogenicity factors, 187–188 Paroxysmal nocturnal hemoglobinuria, see PNH Pathogenesis, PNH, immune model, 120–127 Pathogenicity factors, parasite GPIs as, 187–188

277

Index

Pathophysiology, marrow failure in PNH, 115–120 Phenotypic mosaicism, PNH erythrocytes, 64–68, 88 Phospholipase C GPI-specific, and variant surface glycoprotein, 242–243 mediated cleavage of GPI-anchored proteins, 259 phosphatidylinositol-specific, 201–202, 240–242 Phospholipase D, GPI-specific, 244–245, see also GPI-PLD Physicochemistry, detergent insolubility, 226–227 Physiological role, GPI-PLD, 257–262 Piga characteristics, 141 conditional disruption by means of Cre/ loxP system, 151–153 disruption, and clonal dominance, 150–151 gene inactivation in ES cells, 142–143 PIG-A characterization, 23–24 gene dysfunction, 131 PIG-A(⫺) ES cells, in vitro differentiation, 143–149 PIG-A(⫺) HSC, chimeric mice bearing, 149–154 PIG-A mutation comparison with mutations in FVIII and G6PD, 42–43 correction, 86 detection, 27–31 germline, 32 and GlcNAc-PI synthase defect, 203, 205 and GPI-AP-deficient phenotype, 113–114 in HSC pool, and pathogenesis of PNH, 120–127 in PNH patients, 14–15 somatic recurrent, 40 spectrum, 33 without PNH, 40–41 PIG-A protein part of enzyme complex, 24 structure–function relationships, 41 Pigmenturia, due to destruction of red cells, 2, 4

Pillemer, L. isolation of properdin, 53–54 suggested alternative system of complement activation, 11–12 Plasma acidification, 52 activation of complement alternative pathway, 62–63 and cellular forms of GPI-PLD: relationship, 256–257 Platelets abnormalities in PNH, 104–107 microvesicles, role in PNH, 107 PLD, see Phospholipase D PNH acidified serum test, 50–51 artificial cells, 81–83 clinical behavior in vitro colony culture studies, 116–117 stem cell clonality, 115 therapeutic stem cell transplant, 115–116 diagnosis, 8–15 differentiation of PIG-A(⫺) ES cells in vitro, 143–149 GPI-AP-deficient phenotype, 113–114 hemolysis, treatment, 83–86 historic descriptions, 1–8 incidence of thrombosis, 102–103 patient populations, 101–102 phenotypes and genotypes, correlation, 42 PIG-A mutations, 27–41 platelet abnormalities, 104–107 role of red cell and platelet microvesicles, 107 in somatic cells, biochemical genetics, 21–23 thrombosis in, 12–13 uPAR and cell-mediated fibrinolysis, 108–109 venous thrombosis-associated factors, 103 PNH erythrocytes abnormal sensitivity to complementmediated lysis, 68–83 acidified serum lysis, 53–54 complement sensitivity, 13–14 phenotypes, molecular basis, 77–78 phenotypic mosaicism, 64–68, 88

278

Index

PNH erythrocytes (continued) susceptibility to lysis in acidified serum, 86–87 Point mutations, PIG-A, 33 Polymerase chain reaction, in analysis of PIG-A mutations, 27–30 Prednisone, in treatment of PNH hemolysis, 83–84 Properdin in alternative pathway of complement, 11–12, 64 constitutive presence in serum, 53–54 Protein deficiencies, in PNH, 14 Protein kinase C, activation, GPI role, 186 Proteins apical processing by GPI anchor, 169–170 GPI-anchor addition to, 207–210 Protein truncation test, PIG-A mutations, 30 Protein tyrosine kinase activation, GPI role, 186–187 p56lck, 164 Protozoa GPIs insulin-mimetic activities, 189–190 signal transduction in host cells, 185–187 GPI structures, 180–182 parasitic, GPI-anchored structures, 179–180 R Rafts detergent-resistant membrane associated with, 223–227 enriched in molecules involved in signal transduction, 184–185 GPI-anchored proteins associated with, 160 and GPI-anchor function, 221–222 in living cell membranes, 227–230 role in biosynthetic and endocytic sorting, 230–232 and signaling by GPI-anchored proteins, 232–233 Red cells acid lysis, 5 mechanism of lysis, 8–9 microvesicles, role in PNH, 107

Regulators of complement activation, 58 Resistance, GPI-anchored proteins to degradation by GPI-PLD, 257–258 Respiration, during sleep, effect on hemoglobinemia, 52 RT6, rodent T cell alloantigen, 164–165 Russia, first reported case of PNH, 2 S SCID mouse hematopoiesis by cells from PNH patients, 154–155 human hematopoiesis in, 117–118 Sensitivity PNH erythrocytes to complement, 13–14 PNH erythrocytes to complementmediated lysis, 68–83 residual, GPI-anchored proteins, 257–258 Serotonin, platelet release, effect of complement, 105–106 Short consensus repeats, DAF and CR1, 58 Signal transduction, by protozoal GPIs in host cells, 185–187 Single strand conformation analysis, PIG-A mutations, 28 Sleep, relationship of hemoglobinemia and hemoglobinuria to, 51–53 Somatic cells, PNH, biochemical genetics, 21–23 Sorting, biosynthetic and endocytic, role of rafts, 230–232 Splenectomy, for PNH hemolysis, 84 Splice site mutations, PIG-A, 33 Stem cells clonality, 115 embryonic, see Embryonic stem cells hematopoietic, see Hematopoietic stem cells Strubing, Paul, classic description of PNH, 4–6 Structure–function relationships, PIG-A protein, 41 Survival PIG-A(⫺) HSC, advantage, 155 platelet, in PNH patients, 106–107 Suspension culture, PIG-A(⫺) embryoid bodies grown in, 147

279

Index

T T cell receptor, TCR/CD3 complex, 161–162, 170 T cells CD1d-restricted NK, and immunity to GPI-anchored proteins, 190–191 surface adhesion molecules, GPIanchored ligands for, 125–126 Temperature, cold, and occurrence of PNH, 2, 4 Thrombosis in PNH, 12–13, 102–103 venous, 103 Thy-1, antibody cross-linking, 162 Thymic shared antigen, TSA-1, antibodies, 162 TIMs, see Triton-insoluble membranes Tissues GPI-anchored protein release from, GPI-PLD role, 258–260 GPI-PLD distribution, 252–257 neural, GPI-linked proteins in, 165–166 Toxins, parasite GPIs as, 187–188 Transfusion, saline-washed blood, for PNH hemolysis, 84 Transgenes, hCMV-Cre, 151, 153 Treatment hemolysis of PNH, 83–86 marrow failure in PNH, 127–130 thrombosis in PNH, 109

Triton-insoluble membranes, 211–213 Trypanosoma cruzi, ceramide-containing GPIs, host cell regulation by, 189 Trypanosoma sp. GPI-anchored proteins, 179–180 structures of GPIs, 181 Trypanosomes, variant surface glycoprotein gene expression, 243 U uPAR, see Urokinase–plasminogen activator receptor Urokinase–plasminogen activator receptor and cell-mediated fibrinolysis, 108–109 platelet abnormalities, 104–105 PNH monocytes lacking, 13 various actions, 167 V Variant surface glycoprotein, and GPIspecific phospholipase C, 242–243 Y yGPI8, defective gene, cloning, 206 Young–Luzzatto hypothesis, 15 Z Zinc ions, GPI-PLD dependent on, 245–246 Zymogens, C1r and C1s, 57

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