Current Topics in Cellular Regulation, Volume 35 (Current Topics in Cellular Regulation)

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CURRENT TOPICS IN

Cellular Regulation Volume 35

This Page Intentionally Left Blank

CURRENT TOPICS IN

Cellular Regulation edited by Earl R. Stadtman

National Institutes of Health Bethesda, Maryland

P. Boon Chock

National Institutes of Health Bethesda, Maryland

Volume 35

ACADEMIC PRESS San Diego New York Boston London Sydney Tokyo Toronto

This book is printed on acid-free paper. @ Copyright © 1997 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. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher's consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923) for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-1997 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0070-2137/97 $25.00 Academic Press 15 East 26th Street, 15th floor. New York, New York 10010 http://www.apnet.com Academic Press Limited 24-28 Oval Road, London NWl 7DX, UK http://www.hbuk.co.uk/ap/ International Standard Serial Number: 0070-2137 International Standard Book Number: 0-12-152835-9 PRINTED IN THE UNITED STATES OF AMERICA 97 98 99 00 01 02 EB 9 8 7 6 5 4 3 2

Contents

Regulation of Iron Metabolism in Eukaryotes TRACEY ROUAULT AND RICHARD KLAUSNER I. Need for Regulation of Iron Metabolism II. Translational Regulation of the Iron Sequestration Protein Ferritin III. Expression of TfR Regulated by Intracellular Iron Levels IV. IREs as Binding Sites for IRP V. Steric Hindrance Model to Account for Function of IRPl in Regulation VI. Iron-Sulfur Cluster of IRPl as Key to Sensing of the Iron Levels ... VII. Residues in Active Site Cleft Essential in IRE Binding VIIL Role of IRPl in Vivo IX. Assembly and Disassembly of Iron-Sulfur Cluster of IRPl X. IRP2 as Ubiquitous IRE Binding Protein Regulated by Degradation When Iron Levels Are High XL Cysteines Indispensable to Function of Degradation Domain XII. IRPs Expressed in Cells: Redundant Function or Undefined Unique Roles XIII. Differences in IRE Binding Sites of IRPl and IRP2 Permit IRPs to Bind Unique Targets XTV. Role for IRE Mutations in Human Disease XV. Speculations on Evolution of IRPs XVI. Other G6nes That May Be Regulated by IRE-IRP Regulatory System XVII. Summary and Conclusions References

1 2 3 3 4 5 6 8 8 9 10 11 12 13 13 14 16 16

Structure, Mechanism, and Specificity of Protein-Tyrosine Phosphatases ZHONG-YIN ZHANG I. Introduction II. Structural Characterization

21 25

VI

CONTENTS

III. Catalytic Mechanism IV. Substrate Specificity V. Conclusion and Perspective References

30 48 62 63

Regulation of Fas-Mediated Apoptosis ROBERTA A. GOTTLIEB AND BERNARD M . BABIOR I. II. III. IV.

Introduction Regulatory Factors Regulation of Fas-Mediated Apoptosis Conclusion References

69 72 86 92 92

Aging and Regulation of Apoptosis HuBER R. W A R N E R

I. II. III. IV. V. VI. VII.

Introduction Genes Involved in Apoptosis Cell Senescence and Apoptosis Immunological Aging and Apoptosis Caloric Restriction and Apoptosis Neurodegenerative Disease and Apoptosis Summary References

107 108 110 112 114 115 117 119

Gene Regulation by Reactive Oxygen Species FiLiBERTO CiMiNO, F R A N C A E S P O S I T O , R O S A R I O A M M E N D O L A , A N D

TOMMASO RUSSO I. Introduction II. Sensitivity of Transcription Factors to Intracellular Redox Changes III. Effects of Redox Changes on Regulation of Gene Expression IV. ROS and Extracellular Signal Transduction V. Concluding Remarks References

123 124 134 139 143 144

Regulation of N F - K B and Disease Control: Identification of a Novel Serine Kinase and Thioredoxin as Effectors for Signal Transduction Pathway for N F - K B Activation TAKASHI OKAMOTO, SHINSAKU SAKURADA, JIAN-PING YANG, AND JOCELYN P . M E R I N

I. Introduction II. Transcription Factor NF-KB and Its Activation Pathways

149 150

CONTENTS III. Involvement of N F - K B in Disease Processes IV. Screening Strategy for Anti-NF-zcB Compounds V. Summary References

Vll 154 157 157 158

Regulation of Bacterial Responses to Oxidative Stress JUDAH L . ROSNER AND GiSELA S T O R Z I. Introduction II. Regulators of Escherichia coli Responses to Oxidative Stress III. Oxidative Stress Responses in Salmonella, Haemophilus, Mycobacterium, and Bacillus IV. Concluding Remarks References

163 164 169 174 175

Mechanism and Regulation of Bone Resorption by Osteoclasts NOBUHIKO K A T U N U M A

I. Historical Background and Future Prospects II. Suppression of Bone Resorption by Cathepsin L Family Inhibitors III. Functional Differentiation of Osteoclasts and Macrophages with Respect to Cathepsins L and B IV. Mechanism and Regulation of Procathepsin L Secretion from Osteoclasts V. Cathepsin L Secreted from Osteoclasts as Precursor Form and Processed by Cysteine Proteinase(s) in Bone Lacunae VI. Inhibitory Mechanisms of H*-ATPase, Carbonic Anhydrase II, and Monensin on Pit-Forming Assay VII. Suppression of Bone Resorption by Cathepsin L Family Inhibitors in Vivo VIII. Possible Strategies for New Drug Design to Protect Bone Resorption References INDEX

179 180 183 184 187 188 189 190 191 193


CURRENT TOPICS IN CELLULAR REGULATION, VOLUME 35

Regulation of Iron Metabolism in Eukaiyotes TRACEY ROUAULT RICHARD KLAUSNER

Cell Biology and Metabolism Branch National Institutes of Child Health and Human Disease National Institutes of Health Bethesda, Maryland 20892

I. Need for Regulation of Iron Metabolism Iron is indispensable to the function of many proteins in prokaryotes and eukaryotes, including enzymes t h a t are involved in such fundamental processes as respiration, photosynthesis, and nitrogen fixation. Features t h a t account for the frequent use of iron in prosthetic groups include the ability of iron to facilitate oxidation-reduction reactions and the flexible coordination chemistry of iron. Because iron is required for vital functions, systems t h a t are devoted to the efficient solubilization and uptake of iron have evolved in cells. In addition to solving the problem of ensuring t h a t supplies of iron are sufficient, cells must also provide defenses against iron-related toxicity. Free iron can participate in reactions that produce toxic by-products, the most well known of which is the Fenton reaction in which reduced metal species, such as ferrous iron, react with hydrogen peroxide to form superoxide anion and hydroxyl radical. Hydrogen peroxide is formed in cells as a spontaneous and potentially toxic by-product of respiration, and its rapid breakdown into water and oxygen is enzymatically facilitated by cellular catalases to protect against formation of toxic oxygen species. The highly reactive hydroxyl radical formed in the Fenton reaction can abstract electrons from target molecules in the cell, including DNA, proteins, and lipids, thereby producing unstable free radicals and irreversible damage in target molecules (1,2). The potential toxicity of these reactions may explain why proteins t h a t are devoted to intracellular iron sequestration are maintained in many cell types. Through regulation of the processes of both iron uptake and sequestration, eukaryotic cells can maintain tight control over amounts of free iron and avoid iron-related toxicity. In this chapter, we will discuss regulation of iron metabolism in eukaryotic cells, focusing on regulation of the iron sequestration pro-

Z.

TRACEY ROUAULT AND RICHARD KLAUSNER

tein, ferritin, and regulation of the iron uptake receptor, the transferrin receptor (TfR). The expression levels of both ferritin and the TfR are regulated by two proteins known as iron regulatory proteins 1 and 2 (IRPl and IRP2), formerly known as iron-responsive element (IRE) binding proteins (IRE-BPs) (3,4), as iron regulatory factor (IRF) (5), or as ferritin repressor protein (FRF) (6). Discussion of the mechanisms of regulation of these proteins will provide an overview of mechanisms of posttranscriptional gene regulation, as four different types of such regulation are involved at different steps in iron regulation. Ferritin is translationally regulated, whereas the TfR is regulated through regulation of mRNA degradation (7-9). IRPs sense iron levels directly, either through changes in the status of an associated prosthetic group of IRPl or through an iron-dependent change in the rate of degradation of IRP2. Coordination of uptake and sequestration to the needs of the cell is accomplished by the regulatory proteins IRPl and IRP2. II. TranslatJonal Regulation of the Iron Sequestration Protein Ferritin Ferritin is a multimeric protein composed of 24 subunits which coassemble to form a hollow sphere (10). Within the sphere, iron is precipitated as iron hydroxyphosphate. Subunits are of two types: the H chain, which is heavier (approximately 20 kDa) and is the predominant form in heart, and the L chain, which is lighter (approximately 18 kDa) and is the predominant form in liver cells. The functional H chain gene in humans is located on chromosome 11, whereas the functional L chain gene in humans is located on chromosome 19 (11). The 5'UTRs of ferritin transcripts contain cis-acting elements that mediate the translational regulation of ferritin. When cells are iron-replete, mRNAs encoding ferritin subunits are freely translated, whereas when cells are depleted of iron, the translation of these transcripts is repressed. Shorter sequences within the 5'UTRs of ferritins of H and L chains of numerous species responsible for mediating the translational regulation of these transcripts were first identified by deletional analyses of the 5'UTRs (12,13). The functional motifs within 5'UTRs form RNA stem-loop motifs known as iron-responsive elements, or IREs (14) (see Fig. 1). Phylogenetic comparisons, mutational analyses, and comparisons among functional IREs in different species and between H and L chains have permitted definition of the features of a consensus IRE. An IRE consists of a base-paired stem interrupted by an unpaired cytosine five base pairs removed from a 6-membered loop. The sequence of the loop is

REGULATION OF IRON METABOLISM IN EUKARYOTES

6

almost always CAGUG(X), where X can be any base but G (7). The fact that the IRE is sufficient to mediate translational regulation was established by studies of chimeric transcripts. When IREs were positioned in the 5'UTR of reporter genes, the transcripts became translationally regulated by iron (13,14), confirming that the IRE could transfer iron-mediated translational regulation to unrelated mRNAs expressed in mammalian cells.

III. Expression of TfR Regulated by Intracellular Iron Levels When cells are iron-replete, levels of TfR decrease, whereas in cells treated with iron chelators, synthesis and expression of TfRs are high. These changes in levels of protein expression refiect changes in the levels of the mRNA for TfR (7-9). In iron-depleted cells the mRNA for TfR is relatively stable, whereas in iron-replete cells the mRNA is rapidly degraded. Five IREs are found in the 3'UTR of the TfR transcript, and when the IRE-containing portion of the 3'UTR is ligated to reporter genes, the mRNA half-life of those genes becomes regulated by iron levels (5,15). Structural studies using oligonucleotides and RNase mapping have confirmed that IRE structures are present in the 3'UTR of the TfR (16). When IREs are mutated, the TfR transcript is rapidly degraded regardless of the iron status of the cell. There is a rapid turnover determinant in the 3'UTR between IREs which is required for mRNA degradation (17).

IV. IREs as Binding Sites for IRP Gel retardation assays were the initial means by which cytosolic regulatory proteins that bound to IREs were identified (18,19). Lysates were shown to contain an IRE binding protein that bound to IREs with high affinity (K^ = 10-50 pM) and specificity (20,21). The gel retardation assay was used to monitor purification of the IRE binding protein, which was accomplished using RNA affinity chromatography (22), traditional column chromatography (23,24), or a combination of the two approaches (25). A second assay used to monitor purification of the protein binding to IREs was based on repression of translation of ferritin mRNA in vitro, and the trans-acting protein was therefore referred to as the ferritin repressor protein (FRP) (6). In gel retardation assays performed on lysates of cells, IRE binding activity accurately reflects the iron status of the cells prior to lysis. When cells are irondepleted, IRE binding activity is high, whereas in cells that are ironreplete, binding activity is low (20).

4


Cloning of the IRE binding protein (3,23,24,26), now referred to as IRPl, proved to be unexpectedly informative as the sequence of IRPl turned out to be almost 30% identical to that of porcine mitochondrial aconitase (27). Mitochondrial aconitase had been the subject of extensive biochemical studies (28,29) and the crystal structure was known (30,31). The enzymatic active site residues of mitochondrial aconitase had been identified and studied by site-directed mutagenesis (32), and though overall sequence identity between mitochondrial aconitase and IRPl was only 28%, all of the active site residues between the two proteins were identical (27). Mossbauer spectroscopy and biochemical studies had established that mitochondrial aconitase contained an ironsulfur cluster (33,34) which was cubane (35). Of the four irons in the [4Fe-4S] cluster, three were shown to be directly bound to cysteines of the peptide, whereas the fourth had potentially open coordination sites and could bind directly to the enzymatic substrates citrate, isocitrate, and cis-aconitate (31). An analogous cluster was found in IRPl in ironreplete cells, and IRPl was revealed to be the source of aconitase activity in the cytosol of eukaryotes (36).

V. Steric Hindrance Model to Account for Function of IRP1 in Regulation When the IRP is bound to IREs in the 5'UTR of transcripts, the protein interferes with translational initiation and prevents active translation of the mRNA transcripts of ferritin and erythocyte aminolevulinic acid synthase (eALAS, 5-aminolevulinate synthase), the enzyme which is the rate-limiting step in the biosynthesis of heme (3740). When IRP is bound to IREs in the 3'UTR of the transferrin receptor, binding prevents cleavage of the TfR mRNA at an endonucleolytic cleavage site, most likely by inhibiting access of RNases to the site (17). In both cases, it is likely that binding of IRP creates a steric block. For IREs that are located in the 5'UTR, proximity to the 5' cap is required for function, and there is a loss of translational regulation when the IRE is located more than 67 nucleotides 3' of the cap site (41). The assembly of the 43S subunit of the ribosome at the 5' cap is inhibited (42). The mechanism is likely steric blockage of translational initiation as binding of other unrelated proteins to RNA motifs in the 5'UTR can also inhibit translation (43). Similarly, it appears that IRPs bound to IREs in the 3'UTR may prevent access of an endonuclease to a preferred cleavage site (17). Thus, binding by IRPs simultaneously prevents synthesis of ferritin and eALAS and indirectly increases synthesis of the TfR by increasing the amount of TfR transcript (see Fig. 1).

REGULATION OF IRON METABOLISM IN EUKARYOTES Ferritin mRNA

One IRE in 5'UTR AAAAAA

40S c ribosomal subunit

AAAAAA

60S Q 7

IRE is occupied by IRP inhibiting translation initiation.

ribosomal subunit

"^^ '® ""occupied allowing polysome formation and ferritin synthesis.

Five IREs in 3'UTR

TfR mRNA Protein Coding

ftffl^

AAAAAA

ft

One or more IREs are occupied by IRP protecting mRNA from degradation.

ft

/^ AAAAAA

IREs are unoccupied rendering mRNA susceptible to endonuclease which may be rate-determining step in mRNA degradation.

FIG. 1. A model for IRP-mediated regulation of expression of ferritin and TfR. Binding of IRP to the IRE at the 5' end of ferritin mRNA prevents initiation of translation in iron-depleted cells, whereas translation proceeds in the iron-replete cell. Binding of IRP(s) to the IREs in the 3'UTR of the TfR protects an endonucleolytic cleavage site, which is readily cleaved in iron-replete cells in which IRP does not bind IREs.

VI. Iron-Sulfur Cluster of IRP1 as Key to Sensing of Iron Levels Amounts of IRP 1 that are detectable immunologically do not change significantly when cells are made iron-replete or are iron-depleted (44,45). However, changes in intracellular iron status alter the status of the iron-sulfur cluster and the function of the protein. In cells that are depleted of iron, IRPl is found predominantly as an apoprotein.

b


devoid of an iron-sulfur cluster (46-48). The apoprotein form of IRPl binds with high affinity to IREs, whereas the [3Fe-4S] form of the cluster does not bind IREs (46). Studies with recombinant IRPl have shown that the protein functions either as an active cytosolic aconitase containing a [4Fe-4S] cluster in cells that are iron-replete or as a high affinity IRE binding protein in cells that are iron-depleted (47-49). The two functions of IRPl, as cytosolic aconitase (holoprotein) or as the IRE binding protein (apoprotein), are mutually exclusive properties (7,50). When the three cysteines that ligate the iron-sulfur cluster of IRPl are individually mutagenized to serine, the recombinant protein expressed in cells constitutively binds IREs in gel retardation assays of lysates, regardless of the iron status of the cell prior to lysis (51-53). The results of mutagenesis of the cysteines, along with the correlation of aconitase activity with the iron-replete status, supports the view that the iron sulfur cluster is itself the determinant of function. When cells are iron-replete, the holoprotein, which does not bind IREs, is present, whereas when cells are depleted of iron or are incapable of assembling an iron-sulfur cluster because of mutations of cluster ligands, IRPl is found as apoprotein which binds with high affinity to IREs (46,54). VII. Residues in Active Site Cleft Essential in IRE Binding In the crystal structure of mitochondrial aconitase, an active site cleft separates the fourth domain of the protein from domains 1-3. The fourth domain is connected to the first three domains by a hinge-linker peptide which is not highly structured and which would be expected to exhibit considerable fiexibility. Residues that are analogous to the known active site residues of mitochondrial aconitase have been shown to be important in RNA binding. Ultraviolet (UV) cross-linking studies of IRE and protein have identified both short peptide fragments that contain active site residues (55) and longer peptide fragments from the presumed active site cleft of the protein (56,57). Arginine residues from both sides of the putative active site cleft have been shown to be important in high affinity IRE binding (53). These arginine residues align with known enzymatic active site residues of mitochondrial aconitase and are thought to contribute to the enzymatic active site of IRPl. When these arginines are mutagenized to glutamine, aconitase activity of IRPl is eliminated (53). Although the residues identified as being important in IRE binding are believed to be located in the putative active site cleft of IRPl, the dimensions of the cleft, as revealed in the crystal structure of


7

mitochondrial aconitase, are such that an RNA stem-loop would be physically excluded from this area (see Fig. 2) unless the dimensions of the active site cleft were changed. The fact that IRPl functions as a cytosolic aconitase that is just as efficacious as mitochondrial aconitase supports the view that the geometry of the active site is very similar in the two proteins (36). However, there is no information about the geometry of the IRE binding site in the apoprotein, as crystals of the IRE bound to protein have not been obtained. It is likely that the flexible hinge-linker permits movement of the fourth domain of IRPl with respect to the remainder of the protein and that potential binding surfaces that were previously not accessible become contacts for IREs (see Fig. 2). The fact that the RNA binding site overlaps with the enzymatic active site explains why the two activities are mutually

flexible hingelinker structure of mitochondrial aconitase FIG. 2. A model of mitochondrial aconitase showing the flexible hinge-linker which connects domains 1-3 to domain 4. Residues analogous to those that have been implicated in IRE binding are illustrated in black space-fill form. Residues 100 and 101 of IRP were identified as important to IRE binding in UV cross-linking, and the residues that align with those residues are shown in the backbone alignment of IRPl. Residues R780, R541, and R536 of IRPl were identified as important in IRE binding by IRPl and IRP2 (see text), and the corresponding residues of mitochondrial aconitase are labeled in the ribbon diagram of mitochondrial aconitase. The IRE binding affinity of R780Q to IRPl was decreased 10,000-fold; R541Q was decreased 1000-fold; R536Q was decreased 100-fold. DlOO HlOl was contained within cross-linked peptide.

8


exclusive properties of the protein. In mitochondrial aconitase, the substrate binds to residues on either side of the active site cleft that separates domain 4 from the first three domains (31), a feature which would be expected to contribute to the apposition of domain 4 to domains 1-3 in the holoprotein. Although the active site residues identified as important in IRE binding by UV cross-linking and mutagenesis are present in mitochondrial aconitase, mitochondrial aconitase does not bind IREs (58). The aconitase of Escherichia coli is highly related to IRPl, with a 60% sequence identity between the proteins and identity between active site residues (59), and yet there is also no indication that the E. coli aconitase can bind IREs. Clearly, although some important observations have been made on residues that contribute to the IRE binding site, much remains to be learned about the IRE binding site, as it is not clear why neither mitochondrial aconitase nor E. coli aconitase can bind IREs. There is one circumstance in which it appears that holoprotein can bind IREs: when the holoprotein is treated with high concentrations of reducing agents, including 2% 2-mercaptoethanol (v/v), high affinity binding of IREs is induced (60). The cluster does not dissociate completely from the protein, as high concentrations of 2-mercaptoethanol do not convert the protein to the IRE binding form permanently, and the protein ceases to bind IREs as soon as the 2-mercaptoethanol is removed by chromatography. In this situation, it is possible that the cluster is sufficiently altered by the treatment with the 2-mercaptoethanol that local conformation is changed, permitting access to residues that would normally be inaccessible for IRE binding. VIII. Role of IRP1 In Vivo When the form of IRPl which contains cysteine-to-serine mutations is expressed in cells, regulation of ferritin and TfR is impaired. Cells are unable to derepress ferritin translation when iron-replete and are similarly unable to decrease biosynthesis of the TfR in iron-replete cells (61). These studies of the effect of the mutagenized protein on intracellular iron metabolism support the hypothesis that the ironsulfur cluster of IRPl permits sensing of intracellular iron levels and determines regulatory function in vivo. IX. Assembly and Disassembly of Iron-Sulfur Cluster of IRP1 In the regulation of IRPl, it is important to understand the mechanisms of assembly and disassembly of iron-sulfur clusters, as it is


9

apparent that the iron-sulfur cluster is the key determinant of whether the protein will bind to IREs. The iron-sulfur clusters of many proteins are known to be degraded by oxidants (61a), and the key to stability of these prosthetic groups appears to be determined by whether the cluster is positioned in the protein such that it is accessible to oxidants and solvent. In the case of IRPl, the cluster is positioned in a hydrophilic cleft in the protein where it is readily accessible to oxidants and solvent. The iron-sulfur cluster of IRPl is destabilized by exposure to nitric oxide (62-64) or to hydrogen peroxide (65,66). In addition, the ironsulfur cluster of E. coli aconitase, which is 60% identical in primary sequence to IRPl, is known to be rapidly destabilized by superoxide anion (67), and other hydrolases ofE. coli that contain [4Fe-4S] clusters are similarly sensitive to superoxide (68). Thus, it is quite possible that the iron-sulfur cluster of IRPl is relatively easily degraded under conditions of aerobic growth, whereas the protein itself may have a relatively long half-life. In a setting in which the iron-sulfur cluster is turned over much faster than the protein itself, it is possible to imagine how the iron-sulfur cluster could allow the protein to serve as an iron sensor: the cluster would be reassembled when sufficient iron and sulfur were present with the aid of cluster assembly enzymes, but if iron levels were low, the cluster would not be reassembled and apoprotein, the form which represses translation of ferritin and increases biosynthesis of the TfR, would accumulate (61a). X. IRP2 as Ubiquitous IRE Binding Protein Regulated by Degradation When Iron Levels Are High At the time of cloning of IRPl, the cDNA encoding a second, highly related protein, IRP2, was cloned (3). The protein was 58% identical in primary amino acid sequence, and recombinant protein expressed from the sequence was found to bind consensus IREs with high affinity similar to IRPl (4). However, there were several noteworthy differences between IRPl and IRP2. Unlike IRPl, the predicted active site residues of IRP2 were not identical to those of mitochondrial aconitase; in particular, the serine proposed to function as the catalytic base in the aconitase reaction was noted to be a glutamine at the analogous position in IRP2. In mitochondrial aconitase, mutation of the active site residue results in a loss of aconitase activity (32), and there is no detectable aconitase activity associated with IRP2 (69,70). In addition to the lack of aconitase activity, IRP2 differs from IRPl in the mode of regulation by iron. Unlike IRPl, which remains present in the iron-replete cell, although the IRE binding activity is lost, IRP2 is physically absent in iron-replete cells (69,70). Experiments have

10


shown t h a t the rates of synthesis of IRP2 are equal in iron-replete and iron-depleted cells but t h a t the rates of degradation vary markedly (70,71). In the iron-replete cell, IRP2 is degraded within minutes to hours, whereas in the iron-depleted cell, IRP2 is relatively stable, with a half-life of 9-12 h r (45,72,73). One notable difference between the two proteins is t h a t IRP2 contains an insertion of 73 amino acids relative to I R P l , and the amino acid insertion is encoded by a separate genomic exon. When the amino acids corresponding to the unique exon are excised, the remaining protein is still able to bind IREs with high affinity and is therefore not misfolded as a result of the deletion. However, rapid degradation in the presence of iron is no longer seen. When the IRP2-specific exon is ligated to the analogous position in an expression construct of I R P l , the chimeric I R P l acquires the ability to be rapidly degraded in the presence of iron (72). Thus, the IRP2-specific exon is an iron-dependent degradation domain, capable of conferring the rapid degradation phenotype on a recipient protein t h a t is otherwise stable in iron-replete cells.

XI. Cysteines Indispensable to Function of Degradation Domain The degradation domain of IRP2 has an unusual distribution of amino acids, containing 5 cysteines, 10 prolines, and 8 glycines. When 3 of the cysteines are mutagenized simultaneously to serines, the rapid degradation in iron-replete cells is no longer seen (72). It is quite likely t h a t the cysteines directly bind iron, either singly or as part of an ironsulfur cluster. Although there is still no direct proof t h a t iron is bound by the degradation domain, it is attractive to hypothesize t h a t direct binding of iron to the degradation domain leads to a change in the protein t h a t results in targeting for degradation. One possibility is t h a t a conformational change driven by the binding of iron results in surface exposure of an epitope t h a t is recognized as a degradation target. Another is t h a t ferrous iron bound but not fully coordinated by the protein can react with hydrogen peroxide to release hydroxyl radicals t h a t cause damage in the vicinity of the iron-binding site. A similar process, known as metal-catalyzed oxidation, has been described for the degradation of the E, coli protein glutamine synthetase (2) ( g l u t a m a t e ammonia ligase), and is believed to account for turnover of other proteins. IRP2 is degraded by the proteasome, as evidenced by studies which show t h a t inhibitors of proteases, including MG132 and lactacystin, inhibit the degradation of IRP2 (71,72). It is not known whether IRP2


11

is ubiquitinated, and little is known about the process by which IRP2 is targeted to the proteasome.

XII. IRPs Expressed in Cells: Redundant Function or Undefined Unique Roles In cells that have been examined to date, both IRPl and IRP2 are expressed, although the ratios of the two proteins vary among cell types. In most cells with the exception of brain tissue, IRPl is far more abundant than IRP2, as judged by Northern analyses (70) and by gel retardation assays (74) in which the complexes arising from binding of each protein are separable. Because both proteins bind to consensus IREs with equally high affinity, the advantages of expressing both IRPs in individual cells are not obvious. When transcripts containing IREs are translated in in vitro translation systems, IRPl and IRP2 are just as efficacious as translational repressors (75). One obvious difference between the two proteins is that IRPl functions as a cytosolic aconitase in iron-replete cells, whereas IRP2 does not have aconitase activity and is rapidly degraded in iron-replete cells. It is not clear what role cytosolic aconitase has in cytosolic metabolism. Citrate and isocitrate are both present in the cytosol, and citrate gives rise to precursors of fatty acid synthesis, whereas isocitrate is a precursor of glutamate and glutamine synthesis. When cytosolic aconitase is absent in cells, the interconversion of citrate and isocitrate can be performed in mitochondria by mitochondria aconitase, and precursors and products can cross the mitochondrial membrane. There are several approaches that can aid in the determination of the role of each IRP in the cell. They include expression of mutant forms of the protein, as in the case of IRPl, in which a cysteinyl cluster ligand has been mutagenized, or "knockout" of the function of the two IRPs, either individually or simultaneously. When the constitutive IRE binding form of IRPl is expressed in cells, derepression of ferritin synthesis does not take place in iron-replete cells and regulation of the TfR is similarly impaired (61), indicating that IRPl can mediate regulation of these transcripts in vivo. When IRPl function is eliminated in mice through homologous recombination, there is no apparent impairment in regulation of iron metabolism in the mice, and there is no apparent phenotype associated with the loss of cytosolic aconitase activity (K. Iwai, R. D. Klausner, and T. A. Renault, unpublished observations). Elimination of IRP2 activity by homologous recombination should help to establish whether one IRP has a predominant role in iron regulation or whether each IRP serves as a backup for the other.

12


Regulation of protein function is commonly accomplished through degradation of the unwanted activity, as exemplified by regulation of IRP2. Degradation ensures that the unwanted activity cannot be inappropriately activated under certain circumstances, leading to misregulation. In the case of IRPl, it appears that binding activity is activated not only by iron deprivation but also by oxidative stress (see Section IX). In fact, the sensitivity to oxidants may be the means by which the cluster is normally turned over, opening the possibility for sensing of iron levels by the apoprotein. It is quite possible that an important role of IRPl will turn out to be its role as a sensor of oxidative stress, in addition to its role in regulation of iron metabolism.

XIII. Differences in IRE Binding Sites of IRP1 and IRP2 Permit IRPs to Bind Unique Targets Although both IRPl and IRP2 bind IREs with high affinity, each IRP has, in addition, a ligand identified which is bound by one of the two IRPs but not the other. These ligands were identified by performing SELEX procedures on IRPl (76,77) and IRP2 (77). SELEX, which stands for selective evolution of ligands by exponential enrichment (78), is a procedure in which the protein selects from a group of randomized RNA ligands those ligands which are bound with the highest affinity. The results of the studies performed with IRPl and IRP2 have shown that each IRP has, in addition to the ability to bind the consensus IRE, specificity for a ligand that is not bound by the other IRP. In IRPl, the alternative ligand for IRPl has the sequence UAGUA(X) in the loop (76,77). Since both the consensus IRE and the U1A5 alternative ligand have the potential to form base pairs between the first and fifth nucleotides of the loop, it is possible that base pairing in the loop is present in the bound IRE. Such issues could be resolved by cocrystal structures. IRP2 binds a ligand with the sequence GGGAG(X) where the sixth nucleotide can be any one of the four nucleotides. This sequence is bound with lower affinity than the consensus IRE and does not appear to be capable of mediating translational regulation in an in vitro translation assay (77). The GGGAG(X) alternative ligand is bound poorly by IRPl, an observation which emphasizes that the IRE binding sites are different, although they are similar in their ability to bind the consensus IRE. Although endogenous transcripts that contain these alternative ligands have not been found, it is possible that there is a set of endogenous transcripts that is bound by one IRP but not the other. This feature could be valuable if, for instance, IRPl was serving as a sensor of oxidative stress in cells. Under these circumstances.


13

expression of genes that would exacerbate the toxicity of oxidative stress could be translationally repressed by IRPl, and gene products involved in mitigation of oxidative stress could contain alternative ligands in the 3' end which might lead to increased half-life of the mRNA and increased expression. Differences in the RNA binding sites of the two IRPs revealed by differences in ligand specificities are further underscored by sitedirected mutagenesis studies. Whereas mutation of a single arginine of the enzymatic active site of IRPl results in substantial loss of IRE binding affinity, mutation of the analogous arginine in IRP2 has no measurable impact on IRE binding affinity. However, when this mutation is combined with mutations in other active site arginine residues, binding affinity is profoundly decreased, revealing that although similar residues contribute to binding, their relative importance between the two proteins differs (77). XIV. Role for IRE Mutations in Human Disease A new genetic disorder has been described in humans: the hereditary hyperferritinemia-cataract syndrome. Affected individuals have a combination of elevated serum ferritin and congenital bilateral nuclear cataracts (79,80). Mutations have been identified in the IRE of the ferritin L chain; in one case, the mutation changes the sequence of the IRE loop from CAGUGU to CGGUGU, whereas in the other case, the mutation is also in the loop but the sequence is CACUCU. In both cases, the mutations to the loop result in loss of IRP binding, as would be expected on the basis of previous mutagenesis studies (76,81), which have shown that the sequence CAGUG is required for high affinity binding. In a SELEX study of both IRPl and IRP2, the two nucleotides of the consensus sequence that were never substituted in high affinity binding forms were the unpaired bulge C of the stem and the first G of the loop (77), which is mutated in one of the affected families (79). The pathophysiology of hyperferritinemia is clear, but the reason for an association with formation of cataracts is unclear. XV. Speculations on Evolution of IRPs Iron-sulfur clusters may have been one of the earliest types of protein prosthetic groups in widespread use. When early life forms began to evolve, the atmosphere of the Earth was anaerobic, and iron was probably highly abundant and soluble. Geothermal vents may have led to high concentrations of hydrogen sulfide in certain locales. Iron-sulfur

14


clusters of various stoichiometries, including cubane [4Fe-4S] clusters, have been shown to form spontaneously (82) when a reducing atmosphere and high concentrations of iron and sulfide are found (83). Favorable synthetic conditions may have been present in certain locales early in the history of the Earth, and this could explain why iron-sulfur clusters are found as prosthetic groups in many enzymes central to metabolism, including several enzymes of the Krebs cycle, a central metabolic pathway in which precursors for other metabolic pathways are synthesized. The ancestral protein to I R P l was probably the aconitase ofE. coli, which is 60% identical to I R P l (59), a homology which is striking and which far exceeds the homology between the mammalian mitochondrial and cytosolic aconitases. Much experimental evidence supports the view t h a t iron-sulfur clusters are quite labile in the presence of oxidants (61a). If the iron-sulfur cluster of aconitase is accessible to solvent and oxidants in solution, it could be degraded and subsequently regenerated by iron-sulfur cluster assembly processes in the cell. However, it seems apparent t h a t such regeneration of the cluster could take place only in the presence of sufficient iron and sulfur, and t h a t apoprotein would accumulate in iron-depleted cells. On the basis of the relationship of I R P l to mitochondrial aconitase, we have proposed t h a t the loss of the iron-sulfur cluster permits motion in the region of a flexible hinge-linker of the fourth domain with respect to domains 1-3 and thereby permits access of large molecules to a portion of the protein which is normally accessible only to solvent. Under these conditions, binding of endogenous mRNAs to sequence spaces specific to the apoprotein could occur. Because the binding site would be available only in iron-depleted cells, it would be reasonable that such a site could be incorporated into regulatory pathways.

XVI. Other Genes That May Be Regulated by IRE-IRP Regulatory System Other pathways t h a t could benefit from regulation by an IRE binding protein could include pathways in which proteins require incorporation of iron for function. In the interests of economy, synthesis of such proteins could be repressed during times of relative iron depletion, since the product of synthesis would not fulfill its physiological role without iron. An example of such economy exists with the example of the eALAS protein, the enzyme proposed to be the rate-limiting step in the biosynthesis of heme. Heme is the product of incorporation of iron into protoporphyrin IX, and it is not only nonfunctional in the


15

absence of iron, but there is significant toxicity associated with the accumulation of heme precursors. An IRE present in the 5'UTR of eALAS prevents active biosynthesis of ALAS in erythrocyte cells t h a t are iron-depleted, and the resulting economies to the cell are clear (39,40). It is interesting to note t h a t IREs are contained in two enzymes of the Krebs cycle, succinate dehydrogenase subunit b (SDHb) ofDrosophila melanogaster (84) and porcine mitochondrial aconitase (38,85). The transcripts of four different mammalian species—human, porcine, bovine, and mouse—contain identically conserved IREs in the 5'UTR of mitochondrial aconitase, and this conservation among species in the 5'UTR, which is normally quite divergent among species, implies a potentially important role in regulation of this protein and in physiology. In fact, it appears t h a t this IRE is sufficient to mediate translational regulation in in vitro translation systems; furthermore, there are demonstrable differences in total levels of mitochondrial aconitase in tissues of animals on high versus low iron diets (86). In the case of the IRE found in succinate dehydrogenase (SDHb) of Drosophila, the IRE is sufficient to mediate translational regulation of a chimeric gene expressed in mammalian cells and to shift the distribution of SDHb mRNA from polysomes (actively translated) to mRNPs (translationally repressed) when the Drosophila cells are treated with iron chelators (87). Interestingly, an IRE is not found in h u m a n SDHb mRNA (88), but the h u m a n mitochondrial aconitase gene contains a consensus IRE (86). It is interesting to note t h a t these are the two enzymes of the Krebs cycle t h a t require iron-sulfur prosthetic groups. Iron-sulfur clusters are also found in many of the proteins of the mitochondrial electron transport chain. If synthesis of an iron-requiring enzyme of the Krebs cycle is repressed by lack of iron, this could lead to impairment of the function of the entire cycle. Thus, it is possible t h a t the Krebs cycle itself is regulated by iron levels, and therefore t h a t fundamental features of respiration and energy storage are regulated by iron. If the Krebs cycle were made less efficient, oxygen levels would be expected to rise, since oxygen would no longer be as rapidly consumed. The increases in oxygen tension could lead to more rapid oxidative disassembly of the clusters of remaining cytosolic aconitase, which would then lead to increased availability of iron for other cellular processes. If this were true, then the fates of iron and oxygen would be intertwined under conditions of iron excess, in which the reaction of iron with oxygen promotes toxicity, and iron depletion, in which an increase in oxygen tension could lead to an increase in levels of chelatable iron, and thus produce a self-correction in the system.

16


XVII. Summary and Conclusions Iron metabolism is regulated in cells to ensure that iron supplies are adequate and nontoxic. The expression of iron metabolism is regulated primarily by posttranscriptional mechanisms. Ferritin, eALAS, SDHb of Drosophila, and mammalian mitochondrial aconitase are translationally regulated. The TfR is regulated at the level of mRNA stability. Iron regulatory proteins are regulated either by assembly or by disassembly of an iron-sulfur cluster (IRPl) or by rapid degradation in the presence of iron (IRP2). The list of targets for IRP-mediated regulation is growing longer, and a range of possibilities for versatile regulation exists, as each IRP can bind to unique targets that differ from the consensus IRE. The reactivity of iron with oxygen and the creation of toxic by-products may be the evolutionary stimulus that produced this system of tight posttranscriptional gene regulation. REFERENCES 1. Imlay, J. A., and Linn, S. (1988). Science 240, 1302-1319. 2. Stadtman, E. R. (1993). Annu. Rev. Biochem. 62, 797-821. 3. Rouault, T. A., Tang, C. K , Kaptain, S., Burgess, W. H., Haile, D. J., Samaniego, F., McBride, O. W., Harford, J. B., and Klausner, R. D. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 7958-7962. 4. Rouault, T. A., Haile, D. H., Downey, W. E., Philpott, C. C , Tang, C , Samaniego, F., Chin, J., Paul, L, Orloff, D., Harford, J. B., and Klausner, R. D. (1992). BioMetals 5, 131-140. 5. MuUner, E. W., Neupert, B., and Kuhn, L. C. (1989). Cell (Cambridge, Mass.) 58, 373-382. 6. Brown, P. H., Daniels-McQueen, S., Walden, W. E., Patino, M. M., Gaffield, L., Bielser, D., and Thach, R. E. (1989). J. Biol. Chem. 264, 13383-13386. 7. Klausner, R. D., Rouault, T. A., and Harford, J. B. (1993). Cell {Cambridge, Mass.) 72,19-28. 8. Melefors, O., and Hentze, M. W. (1993). Blood Rev. 7, 251-258. 9. Kuhn, L. C. (1994). Bailliere's Clin. Haematol. 7, 763-785. 10. Theil, E. C. (1987). Annu. Rev. Biochem. 56, 289-315. 11. Worwood, M., Brook, J. D., Cragg, S. J., Hellkuhl, B., Jones, B. M., Perara, P., Roberts, S. H., and Shaw, D. (1985). J. Hum. Genet. 69, 159-164. 12. Hentze, M. W., Rouault, T. A., Caughman, S. W., Dancis, A., Harford, J. B., and Klausner, R. D. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 6730-6734. 13. Aziz, N., and Munro, H. N. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 8478-8482. 14. Hentze, M. W., Caughman, S. W., Rouault, T. A., Barriocanal, J. G., Dancis, A., Harford, J. B., and Klausner, R. D. (1987). Science 238, 1570-1573. 15. Casey, J. L., Koeller, D. M., Ramin, V. C , Klausner, R. D., and Harford, J. B. (1989). EMBO J. 8, 3693-3699. 16. Horowitz, J. A., and Harford, J. B. (1992). New Biol. 4, 330-338. 17. Binder, R., Horowitz, J. A., BasiHon, J. P., Koeller, D. M., Klausner, R. D., and Harford, J. B. (1994). EMBO J. 13, 1969-1980. 18. Leibold, E. A., and Munro, H. N. (1988). Proc. Natl. Acad. Sci. U.S.A. 85,2171-2175.


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48. Gray, N. K , Quick, S., Goossen, B., Constable, A., Hirling, H., Kuhn, L. C , and Hentze, M. W. (1993). Eur. J. Biochem. 218, 657-667. 49. Haile, D. J., Rouault, T. A., Tang, C. K., Chin, J., Harford, J. B., and Klausner, R. D. (1992). Proc. Natl. Acad. Sci. U.S.A. 89, 7536-7540. 50. Klausner, R. D., and Rouault, T. A. (1993). Mol. Biol. Cell 4, 1-5. 51. Philpott, C. C , Haile, D. J., Rouault, T. A., and Klausner, R. D. (1993). J. Biol. Chem. 268, 17655-17658. 52. Hirling, H., Henderson, B. R., and Kuhn, L. C. (1994). EMBO J. 13, 4 5 3 - 4 6 1 . 53. Philpott, C. C , Klausner, R. D., and Rouault, T. A., Proc. Natl. Acad. Sci. U.S.A. 91, 7321-7325. 54. Basilion, J. P., Kennedy, M. C , Beinert, H., Massinople, C. M. Klausner, R. D., and Rouault, T. A. (1994). Arch. Biochem. Biophys. 311, 517-522. 55. Basilion, J. P., Rouault, T. A., Massinople, C. M., Klausner, R. D., and Burgess, W. H. (1994). Proc. Natl. Acad. Sci. U.S.A. 9 1 , 574-578. 56. Swenson, G. R., and Walden, W. E. (1994). Nucleic Acids Res. 22, 2627-2633. 57. Neupert, B., Menotti, E., and Kuhn, L. C. (1995). Nucleic Acids Res. 23, 2579-2583. 58. Kaptain, S., Downey, W. E., Tang, C , Philpott, C. C , Haile, D. J., Orloff, D. G., Harford, J. B., Rouault, T. A., and Klausner, R. D. (1991). Proc. Natl. Acad. Sci. U.S.A. 88, 10109-10113. 59. Prodromou, C , Artymiuk, P. J., and Guest, J. R. (1992). Eur. J. Biochem. 204, 599-609. 60. Hentze, M. W., Rouault, T. A., Harford, J. B., and Klausner, R. D. (1989). Science 244, 357-359. 61. DeRusso, P. A., Philpott, C. C , Iwai, K., Mostowski, H. S., Klausner, R. D., and Rouault, T. A. (1995). J. Biol. Chem. 270, 15451-15454. 61a. Rouault, T. A., and Klausner, R. D. (1996). Trends Biochem. Sci. 21, 174-177. 62. Weiss, G., Goossen, B., Doppler, W., Fuchs, D., Pantopoulos, K., Werner-Felmayer, G., Wachter, H., and Hentze, M. W. (1993). EMBO J. 12, 3651-3657. 63. Drapier, J. C , Hirling, H., Wietzerbin, J., Kaldy, P., and Kuhn, L. C. (1993). EMBO J. 12, 3643-3649. 64. Pantopoulos, K., and Hentze, M. W. (1995). Proc. Natl. Acad. Sci. U.S.A. 92, 1 2 6 7 1271. 65. Pantopoulos, K., and Hentze, M. W. (1995). EMBO J. 14, 2917-2924. 66. Martins, E. A., Robalinho, R. L., and Meneghini, R. (1995). Arch. Biochem. Biophys. 316, 128-134. 67. Gardner, P. R., and Fridovich, I. (1991). J. Biol. Chem. 266, 1478-1483. 68. Flint, D. H., Tuminello, J. F., and Emptage, M. H. (1993). J. Biol. Chem. 2 6 8 , 2 2 3 6 9 22376. 69. Guo, B., Yu, Y., and Leibold, E. A. (1994). J. Biol. Chem. 269, 24252-24260. 70. Samaniego, F., Chin, J., Iwai, K , Rouault, T. A., and Klausner, R. D. (1994). J. Biol. Chem. 269, 30904-30910. 71. Guo, B., Phillips, J. D;, Yu, Y., and Leibold, E. A. (1995). J. Biol. Chem. 270, 2 1 6 4 5 21651. 72. Iwai, K , Klausner, R. D., and Rouault, T. A. (1995). EMBO J. 14, 5350-5357. 73. Pantopoulos, K., Gray, N. K , and Hentze, M. W. (1995). RNA 1, 155-163. 74. Henderson, B. R., Seiser, C , and Kuhn, L. C. (1993). J. Biol. Chem. 268,27327-27334. 75. Kim, H. Y., Klausner, R. D., and Rouault, T. A. (1995). J. Biol. Chem. 270,4983-4986. 76. Henderson, B. R., Menotti, E., Bonnard, C , and Kuhn, L. C. (1994). J. Biol. Chem. 269, 17481-17489.


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77. Butt, J., Kim, H., Basilion, J. P., Cohen, S., Iwai, K., Philpott, C. C, Altschul, S., Klausner, R. D., and Rouault, T. A. (1996). Proc. Natl Acad. Sci. U.S.A. 93, 43454349. 78. Tuerk, C, and Gold, L. (1990). Science 249, 505-510. 79. Girelli, D., Corrocher, R., Bisceglia, L., Olivieri, O., De Franceschi, L., Zelante, L., and Gasparini, P. (1995). Blood 86, 4050-4053. 80. Beaumont, C., Leneuve, P., Devaux, I., Scoazec, J. Y., Berthier, M., Loiseau, M. N., Grandchamp, B., and Bonneau, D. (1995). Nat. Genet. 11, 444-446. 81. Jaffrey, S. R., Haile, D. J., Klausner, R. D., and Harford, J. B. (1993). Nucleic Acids Res. 21, 4627-4631. 82. Berg, J. M., and Holm, R. H. (1981). In "Metal Ions in Biology" (T. G. Spiro, ed.), pp. 1-66. Wiley, New York. 83. Maden, B. E. H. (1995). Trends Biochem. Sci. 20, 337-341. 84. Au, H. C., and Scheffler, I. E. (1994). Gene 149, 261-265. 85. Zheng, L., Andrews, P. C., Hermodson, M. A., Dixon, J. E., and Zalkin, H. (1990). J. Biol. Chem. 265, 2814-2821. 86. Kim, H. Y., LaVaute, T., Iwai, K, Klausner, R. D., Rouault, T. A. (1996). J. Biol. Chem. 271, 24226-24230. 87. Kohler, S. A., Henderson, B. R., and Kuhn, L. C. (1995). J. Biol. Chem. 270, 3078130786. 88. Au, H. C., Ream-Robinson, D., Bellew, L. A., Broomfield, P. L., Saghbini, M., and Scheffler, I. E. (1995). Gene 159, 249-253.


CURRENT TOPICS IN CELLULAR REGULATION, VOLUME 35

Structure, Mechanism, and Specificity of Protein-Tyrosine Phosphatases ZHONG-YIN ZHANG

Department ofMolecular Pharmacology Albert Einstein College of Medicine Bronx, New York 10461

I. Introduction Protein tyrosine phosphorylation and dephosphorylation are fundamental cellular signaling mechanisms t h a t control cell growth and differentiation, mitogenesis, cell cycle, metabolism, gene transcription, cytoskeletal integrity, neuronal development, and the immune response (Yarden and Ullrich, 1988; Bishop, 1991; Cantley et al, 1991; Hunter, 1995). Hundreds of protein kinases and protein phosphatases and their substrates are integrated vîthin an elaborate signal transducing network. The defective or inappropriate operation of this network is at the root of such widespread diseases as cancers, diabetes, and autoimmune disorders. Consequently, the characterization of the individual components and the delineation of the circuitry of this regulatory network have emerged as one of the most active fields in biomedical research. In vivo, the level of tyrosine phosphorylation in a given protein is regulated by the opposing actions of protein-tyrosine kinases (PTKs, EC 2.7.1.112) and protein-tyrosine phosphatases (FTPases, EC 3.1.3.48). PTKs are enzymes t h a t catalyze the transfer of the y-phosphate of ATP to the 4-hydroxyl of tyrosyl residues within specific protein/peptide substrates. PTPases are hydrolytic enzymes t h a t remove phosphate from the phosphorylated tyrosine residue(s). The sequence context surrounding the target tyrosine plays a key role in determining its recognition by PTKs and PTPases. Comprehension of the physiological roles of protein tyrosine phosphorylation, and its potential as a mechanism for reversible modulation of protein function and cell physiology, must necessarily encompass the characterization of PTPases in addition to the PTKs. The structure and function of PTKs have been extensively studied (Hunter, 1987, 1991). However, only recently has attention been focused on PTPases. Several reviews of PTPases have appeared 21

Copyright © 1997 by Academic Press. All rights of reproduction in any form reserved. 0070-2137/97 $25.00

22

ZHONG-YIN ZHANG

(Fischer et al, 1991; Charbonneau and Tonks, 1992; Brautigan, 1992; Walton and Dixon, 1993; Zhang and Dixon, 1994; Barford et al, 1995). This chapter will focus primarily on the structure, mechanism, and substrate specificity of the PTPases. A. Classification of PTPases

The PTPases constitute a growing family of enzymes that rival the PTKs in terms of structural diversity and complexity. Unlike protein kinases, where tyrosine-specific and serine/threonine-specific kinases share sequence identity, the PTPases show no sequence similarity with serine/threonine phosphatases or with the broad-specificity phosphatases such as acid or alkaline phosphatases. Collectively, PTPases can be broadly categorized into two groups: receptor-like and intracellular PTPases (Fig. 1). The receptor-like PTPases, exemplified by the leukocyte phosphatase, CD45, generally have an extracellular domain, a single transmembrane region, and one or two C3ôplasmic PTPase domains. The intracellular PTPases, exemplified by PTPIB and the Yersinia PTPase, contain a single catalytic domain and various amino- or carboxyl-terminal extensions, including SH2 domains that may have targeting or regulatory functions. Although many PTPases are proteins with more than 400 amino acids, their catalytic domains are usually contained within a span of 250 residues referred to as the PTPase domain. This domain is the only structural element that has amino acid sequence identity among all PTPases from bacteria to mammals (Zhang e^ al., 1994a). The unique feature that defines the whole PTPase family is the active site sequence (HA^)C(X)5R(S/T) called the PTPase signature motif in the catalytic domain (Zhang et al., 1994d). Interestingly, the PTPase signature motif can also be found in the structures of two additional groups of enzymes that can bring about phosphomonoester hydrolysis, namely, the VHl(VaccmJa open reading frame iifi)-like dual-specificity phosphatases (Guan et al., 1991) and the low molecular weight (18-kDa) PTPases (Zhang and Van Etten, 1990; Cirri et al, 1993) (Fig. 1). The low molecular weight (Mr) and the VHl-like phosphatases display little amino acid sequence identity with the classical PTPases. The only similarities among these three groups of phosphatases are the relative placement of the essential cysteine and arginine residues in the active sites that constitute the PTPase signature motif (HA^)C(X)5R(S/T). Unlike the PTPases, which show substrate specificity strictly restricted to phosphotyrosyl proteins (Sparks and Brautigan, 1985; Tonks et al., 1988; Guan and Dixon, 1990), the dual-specificity phosphatases and the low M^ phosphatases are unusual in that they can utilize protein substrates containing phosphotyrosine, phosphoserine, and phosphothreonine (Gautier et al.,

23

PROTEIN-TYEOSINE PHOSPHATASES

Protein—Tyrosine

I

Phosphatases

-Il"- "I

IÎ

Yop51 PTP1B SHPTP CD45 PTPa

^ ^1^^^ ^1.^^^ ^ ^1^^^ ^ 1 ^ ^ ^ ^

Dual— Specificity

Phosphatases VHR VH1 MKP1 cdc25

Stpl

WvWWWWN

•

PTPase Signature (H/V)C(X)5R(S/T)

^

PTPase

W

Dual—Specificity Phosphatase Catalytic Domain

I I N- or

Catalytic

C-terminal

Motif: Domain

Extension

• M ^ ^

SH2

Domain

Transmembrane Low

Mr

Fibronectin

Region

Phosphatase Type

III

Domain

FIG. 1. Structural features of receptor-like and intracellular phosphatases containing the PTPase signature motif: C(x)5R(S/T).

1991; Guan et al, 1991; Ishibashi et al, 1992; Zhang et al, 1995a). However, despite the variation in the primary structures and the differences in the active site substrate specificities, the PTPases, the dualspecificity phosphatases, and the low Mr phosphatases utilize a common mechanism to effect catalysis (see Section III). B. Biological Functions of PTPases

The functional role of PTPases in cellular signaling processes is just beginning to be appreciated (Sun and Tonks, 1994; Hunter, 1995). Because deregulated PTKs, such as src, Ick, and neu, can function as

24

ZHONG-YIN ZHANG

dominant oncogenes, it has been assumed that at least some PTPases function as products of tumor suppressor genes. Indeed, mutations in SH-PTPl lead to severe immune dysfunction, giving rise to the motheaten phenotype in mice (Shultz et al., 1993). Thus, SH-PTPl may be an important negative regulator of cytokine signaling; its loss results in sustained tyrosine phosphorylation, with consequent enhanced proliferation (KlingmuUer et al., 1995). The Fas-associated phosphatase (FAP-1) inhibits Fas-generated signals that lead to apoptosis (Sato et al,, 1995). The gene for the receptor-like PTPy has been proposed to be a tumor suppressor gene located on chromosome 3p21, a segment frequently altered in renal and lung carcinomas (LaForgia et al., 1991; Wary et al., 1993). In addition, expression of PTPIB has been shown to block transformation mediated by neu (Brown-Shimer et al., 1992) and to partially revert transformation by src (Woodford-Thomas et al., 1992). On the other hand, there is mounting evidence that some PTPases potentiate rather than antagonize the action of PTKs. This behavior enhances mitogenic signaling, leading to cellular transformation. Thus the receptor PTPase CD45, through its capacity to dephosphorylate and activate the src family of PTKs, is essential for initiating downstream signaling processes in response to stimulation of T- and Bcell receptors (Pingel and Thomas, 1989). SH-PTP2 and its Drosophila homolog corkscrew are positive mediators of growth factor signaling (Perkins e^ aZ., 1992;NoguchieâZ., 1994). The cell cycle regulator cdc25 dephosphorylates Tyr-15 of cdc2, thereby activating the cdc2-cyclin B complex, which, in turn, promotes mitosis (Gould and Nurse, 1989; Millar and Russell, 1992). Strikingly, ectopic expression of PTPce produces a transformed phenotype in rat embryonic fibroblasts (Zheng et al, 1992). Indeed, the plant oncogene rolB, from Agrobacterium rhizogenes, has been shown to encode a protein that displays PTPase activity (Filippini et al., 1996). The importance of PTPases in cellular physiology is further emphasized by the fact that they are often targets for microbial or viral intervention. For instance, the pathogenic bacterium Yersinia encodes a PTPase essential for its virulence (Guan and Dixon, 1990; Bliska et al., 1991), and the Vaccinia virus encodes a dualspecificity phosphatase, VHl (Guan et al., 1991), that is essential for viral transcription and infectivity (Liu et al., 1995). Since the discovery of the Vaccinia VHl phosphatase, a number of additional dual-specificity phosphatases have been identified. The mammalian dual-specificity protein phosphatases have surfaced as key regulators of mitogenic signaling pathways as well as the cell cycle itself(SuneifaZ., 1993; Arocae^ aZ., 1995; Poon and Hunter, 1995;Keyse, 1995). The low M^ PTPases, whose biological function is unknown, were

PROTEIN-TYROSINE PHOSPHATASES

25

previously thought to exist only in mammalian species. Genetic studies of fission yeast (Schizosaccharomyces pombe) with temperature-sensitive mutations of cdc25 (i.e., cdc25-22) have identified a gene that, when overexpressed, rescues cdc25-22 (Mondesert et aL, 1994). This gene, named stpl^ (small tyrosine phosphatase), encodes a protein that is highly similar (42% identical) to the mammalian low M^ PTPases. The low Mr PTPases have been shown to dephosphorylate both phosphotyrosyl and phosphoseryl/threonyl protein substrates (Zhang et al, 1995a). These findings suggest t h a t low My. PTPases may represent a subgroup of dual-specificity phosphatases t h a t may be involved in cell cycle control.

II. Structural Characterization A. PTPases Structures The structures of the PTPase catalytic domains of P T P I B (residues 1-321; Barford etal, 1994) and Yersinia PTPase (residues 163-468; Stuckey et al, 1994) have been determined at 2.8 A and 2.5 A resolution, respectively. Although the amino acid sequence of the Yersinia PTPase is only —20% identical to t h a t of PTPIB, it is clear t h a t the two structures share a common secondary structural scaffold, with close similarity in tertiary structure. P T P I B is composed of a single domain, with 8 a helices and 12 j8 strands. Similarly, the Yersinia PTPase is a single domain protein with 5 a helices and 8 j8 strands (Fig. 2). The central feature of the P T P I B and Yersinia PTPase tertiary folds is a highly twisted, mixed /3 sheet fianked by a helices on both sides. The PTPase active site is located within a crevice on the molecular surface and is contained on (32,138, (33, and pi in Yersinia PTPase (^83, i812, j84, and j811 in PTPIB), as well as on the associated helices and loops t h a t form the core of the PTPase catalytic region (Fig. 2). Residues of the PTPase signature motif in the Yersinia PTPase are located on the loop structure between the ^ t u r n at the COOH terminal of (38 (residues 403-406) and the first t u r n of helix a5 (407-410). The same motif in PTPIB (residues 214-222) is located between the COOH terminus of j812 and the first t u r n of Q;4. This loop is termed the PTPase phosphate binding loop or P-loop (Stuckey et al, 1994). The P-loop includes the invariant residue Cys-403 in the Yersinia PTPase, which is located at the base of the catalytic pocket. The pocket is —10 A deep and —13 A wide and is surrounded by five polypeptide loops or turns (al-j81, a6-a7, j82-a2, a3-i84, and ^4-^87) and helix a 3 . Similarly, the catalytic cleft of PTPIB is surrounded by four loops (cel-jSl, /33-ce2, f311-a3, and a5-a6), j85, and j86. The majority of the invariant amino acid residues conserved

26

ZHONG-YIN ZHANG

FIG. 2. Ribbon diagram of the Yersinia PTPase structure. Reprinted with permission from Nature [J. A. Stuckey et al, Nature (London) 370, 571-575 (1994)]. Copyright 1994 Macmillan Magazines Limited.

in all PTPases from bacteria to mammals (Zhang and Dixon, 1994) are located in and around the enzyme active site (Barford et al., 1994; Stuckey et al, 1994). This suggests that all PTPases are likely to recognize phosphotyrosine in a similar way and have similar catalytic mechanisms. As will be discussed below, some of these residues are important for active site integrity, while others are involved in catalysis. Tungstate is a competitive inhibitor of both the Yersinia PTPase and PTPIB. The crystal structures of both the Yersinia PTPase- (Stuckey et al, 1994) and the PTPlB-tungstate complexes (Barford e^ al, 1994) have also been solved and provide information about the interactions between the enzyme and the phosphate moiety of the substrate. The structures of the PTPase-tungstate complexes show that the tungstate oxygen atom(s) ion pair with the positively charged Arg-409 and Arg221 of the Yersinia PTPase and PTPIB, respectively. The Yersinia PTPase-tungstate structure indicates that there are two hydrogen


27

bonds between the guanidinium group of Arg-409 and two of the tungstate oxygen atoms, a and b (Fig. 3). The tungstate oxygen atoms denoted as a, b, and c are hydrogen bonded to the NH amides of the peptide backbone making up the P-loop. The tungstate oxygen atom denoted as d is projecting out of the active site pocket and interacting with the side chain of the invariant Gln-446 in Yersinia PTPase and Gln-262 in PTPIB. This oxygen atom most hkely corresponds to the oxygen atom present in the substrate scissile phosphate ester bond (Stuckey et al, 1994). Thus, the side chain of R409 acts together with the amides of the P-loop to Ugand the tungstate. The invariant Cys residue (Cys-403 in Yersinia PTPase and Cys-215 in PTPIB) has been shown to be essential for PTPase activity and formation of a covalent phosphoenzyme intermediate. The PTPaseWO4 complexes reveal that the Sy atom of Cys-403 in the Yersinia PTPase is poised 3.6 A from the W atom, while the Sy atom of Cys215 in PTPIB is poised 3.1 A from the W atom. In addition, the sulfur atom is directly opposite the scissile bond oxygen d, and is in a position

FIG. 3. The phosphate binding loop with tungstate bound. Only the peptide backbone from residues 403-409 of the Yersinia PTPase is shown. Tungstate is located in the center of the loop structure. The side chain of Cys-403 corresponds to the active site nucleophile which directly attacks the phosphorus atom in a substrate. The tungstate oxygen atoms, a and b, form two hydrogen bonds with the guanidinium group of Arg409. The tungstate oxygen atoms a, b, and c, also form multiple hydrogen bonds with the NH amides of the peptide backbone making up the phosphate binding loop. The tungstate oxygen atom d is projecting out of the active site and is equivalent to the oxygen present in the scissile ester bond. Reprinted with permission from Z.-Y. Zhang etal, Biochemistry 33,15266-15270 (1994). Copyright 1994 American Chemical Society.

28

ZHONG-YIN ZHANG

for an SN2 nucleophilic attack on a substrate's phosphorus atom (Fig. 3). This is consistent with its role as a nucleophile in the catal3rtic mechanism, as will be discussed below. The apparent thiol pifa of the active site Cys-403 was found to be 4.7 (Zhang and Dixon, 1993). This suggests that Cys-403 exists as a thiolate anion at physiological pH. The crystal structure also provides an explanation for the lowering of the pifa of the invariant Cys residue thiol group from 8.5 in aqueous solution to 4.7 in the PTPase active site. Unlike the cysteine proteinases, which stabilize an active site thiolate anion via an ion pair with a protonated histidine, the PTPases stabilize the active site thiolate by an extensive network of hydrogen bonds radiating out from the Ploop (Barford et aL, 1994; Stuckey et al., 1994). The side chain of the invariant His-402 in the Yersinia PTPase is not involved in ion pair interaction with the side chain of Cys-403 but participates in an array of H bonds that includes the carbonyl of Cys-403, the side chain of His402, the Tyr-301 hydroxyl, and finally, the His-270 side chain. A similar network of H bonds including His-214 is also present in the PTPIB structure. Substitution of the invariant histidine would be expected to disrupt this H-bond network and elevate the apparent piâ of Cys-403. Indeed, H402N and H402A mutations in the Yersinia PTPase display apparent thiol pifa of 6.0 and 7.4, respectively (Zhang and Dixon, 1993). A second array of H bonds from the carbonyls of residues Ala-405 and Gly-406 in Yersinia VTVsiSe (Ala-217 and Gly-218 in PTPIB) mediate interaction with the buried guanidinium group of Arg-440 (Arg-257 in PTPIB) to stabilize the thiolate anion or bound tungstate. Finally, the helix dipole of ab in Yersinia PTPase (a4 in PTPIB) is positioned in such a way that it will also contribute to the stability of the anion. B. Structures of VHR and Low Mr Phosphatase

The VHl-like dual-specificity phosphatases and the low M^ phosphatases display little amino acid sequence identity with the classical PTPases. The only similarities among these three groups of phosphatases are the relative placements of the essential cysteine and arginine residues in the active sites that constitute the PTPase signature motif (HA/^)C(X)5R(S/T) (Fig. 4). While the P-loop structure appears to be conserved in PTPases from bacteria to mammals, it is striking to find that the P-loop of the PTPase also has a structure similar to that of the active site loop of VHR (Yuvaniyama et al, 1996). VHR (for VHlrelated; Ishibashi e^ aZ., 1992), which is believed to be responsible for activation of cdk-cyclin complex(es) at some stage of the cell cycle (Aroca et aZ./1995), has emerged as the prototype for the dual-specificity phosphatases. Not only has this particular phosphatase been overex-

29


Sequence

Phosphatase Yop51 PTPIB CD45

V I H V V H V V H

VHl VHR PAC-1

L V H L V H L V H

Low M^

I

R A G V G S A G I G S A G V G |A A G V N R E G Y S Q A G I S H

FTP

L F V

L G N I C

human cdc25

F V H

E F S S E laG P

FIG. 4. The PTPase signature motif. Reprinted with permission from Z.-Y. Zhang et al, Biochemistry 34, 16389-16396 (1995). Copyright 1995 American Chemical Society.

pressed and purified to homogeneity but, in addition, it has been the subject of several detailed enzymological investigations (Zhou et al,, 1994; Denu et al, 1995a,b; Zhang et al, 1995b). The structure of VHR reveals a general fold that occurs in the Yersinia PTPase and the PTPIB structures, and many of the secondary structural elements of PTPases are also present in VHR. Although VHR is smaller than PTPases, the VHR structure retains the same starting and ending secondary structure elements as the PTPase catalytic domains, suggesting that the VHR structure may define a minimal core structure for both dualspecificity phosphatases and the PTPases (Yuvaniyama et al, 1996). In addition to the general lack of sequence identity between the low Mr phosphatases and the PTPases, the low M^ enzymes are also relatively smaller and contain a PTPase signature motif close to the NH2 terminus of the protein, whereas the signature motif occurs closer to the COOH terminus of the PTPases and the dual-specificity phosphatases. The bovine low M^ PTPase has distinct topologies compared with those of the PTPIB, the Yersinia PTPase, and the VHR (Su et al, 1994; M. Zhang et al, 1994). However, it is interesting that residues of the PTPase signature motif (12-19 in the low M^ PTPase, 403-410 in the Yersinia PTPase, 214-222 in PTPIB, and 123-131 in VHR) form a similar loop structure, termed the phosphate-binding loop, between the ^ turn at the COOH terminus of a j8 strand and the first turn of an a helix (Su et al, 1994; M. Zhang e^ al, 1994; Barford et al, 1994; Stuckey et al, 1994; Yuvaniyama et al, 1996). Furthermore, a structural comparison between the bovine low M^ phosphatase and PTPIB reveals that in addition to the confined similarity in the active sites, the central

30

ZHONG-YIN ZHANG

parallel (3 sheet of low Mj. phosphatase superimposes closely with the four central ^8 strands of PTPIB (Barford et al, 1995). Thus, these different phosphatase structures are striking examples of convergent evolution achieving highly similar active site clefts, and the similarities in the conserved active site motifs may suggest a common mechanism to bring about phosphate monoester hydrolysis in these otherwise very differernt molecules. Indeed, the invariant Cys residue has been shown to be essential for phosphatase activity and formation of a covalent cysteinyl phosphoenzyme intermediate (Zhang, 1990; Guan and Dixon, 1991; Wo et al, 1992; Cho et al, 1992; Zhou e^ al, 1994), whereas the invariant Arg residue in the signature motif has been shown to play an important role in substrate binding and transition state stabilization (Zhang et al, 1994d). These phosphatase structures, taken together with earlier biochemical and more recent mutational studies of active site residues, provide a unique opportunity to identify common mechanistic features associated with this novel family of biological catalysts. In addition, the availability of the structures combined with the ability to mutate specific residues provides an opportunity to test these mechanistic hypotheses and identify key residues for catalysis and substrate/inhibitor recognition. III. Catalytic Mechanism A. An Overview The mechanism of phosphate monoester hydrolysis [Eq. (1)] has been the subject of many investigations (CuUis, 1987; Frey, 1989; Thatcher and Kluger, 1989), ROPOi" + H2O ^ ROH + HOPOi"

(1)

because the hydrolysis of phosphate monoesters is linked to energy metabolism, to metabolic transformation and regulation, and, more recently, to a wide variety of signal transduction pathways. Hydrolysis of phosphate monoesters is a thermodynamically favorable process (AG^ < -2.15 kcal/mol), but in the absence of enzymes, phosphate monoesters are almost kinetically inert (Westheimer, 1987). The means by which their reactivity is enhanced by PTPases presents both an old and a new problem in mechanistic enzymology. A central question concerning the mechanism of phosphatases is whether substitution at phosphorus proceeds by a single-displacement or double-displacement pathway. Single displacement involves the direct transfer of a phosphoryl group from the monoester to water. In


31

the double-displacement mechanism, the phosphoryl group is first transferred to a nucleophilic group of the enzyme, forming a phosphoenzyme intermediate which is then hydrolyzed by water. The kinetic and catalytic mechanisms of both alkaline (Kim and Wyckoff, 1991; Coleman, 1992) and acid phosphatases (Van Etten, 1982) have been demonstrated to involve a phosphoserine (Schwartz and Lipmann, 1961) and a phosphohistidine intermediate (Van Etten and Hickey, 1977), respectively. A carboxyl phosphate intermediate is involved in a sarcoplasmic reticulum ATPase-catalyzed reaction (Degani and Boyer, 1973). Detailed studies of the catalytic mechanism of PTPases are beginning to appear in the literature. B. Novel Thiol Phosphate Covalent Intermediate

The PTPase-catalyzed reaction involves a phosphoenzyme intermediate which can be trapped by addition of a denaturant soon after mixing the enzyme with a ^^P-labeled substrate (Zhang, 1990; Guan and Dixon, 1991; Pot et al, 1991; Wo et aL, 1992; Cho et al, 1992). This suggests that the PTPase-catalyzed hydrolytic reaction is nucleophilic in nature, and involves both the formation and the breakdown of a phosphoenzyme intermediate. An analysis of the chemical stability of the trapped phosphoenzyme intermediate in the PTPase-catalyzed reaction suggests that it has the characteristics of a previously undescribed thiophosphate linkage (-S-POi~) (Zhang, 1990; Guan and Dixon, 1991; Wo et al., 1992). Site-directed mutagenesis experiments show that the invariant Cys residue (e.g., Cys-403 in the Yersinia PTPase and Cys-215 in PTPl) is required for PTPase activity (StreuH et al, 1990; Guan and Dixon, 1990; Gautier et al, 1991; Cirri et al, 1993; Zhou et al, 1994). Replacement of the catalytically essential Cys-215 residue in PTPl with Ser destroys its ability to form a phosphoenzyme intermediate, suggesting that the intermediate is a phosphocysteine (Guan and Dixon, 1991; Cirri et al, 1993; Zhou et al, 1994). In addition, the inactivation of PTPases, the dual specificity, and the low Mj. phosphatases by iodoacetate selectively modifies the active site Cys residues of these phosphatases (Camici et al., 1989; Pot and Dixon, 1992; Zhang and Dixon, 1993; Zhou et al., 1994). The participation of a cysteine residue in the phosphoenzyme intermediate is further supported by ^^P NMR analysis of the trapped intermediate in the low M^ PTPase (Wo et al., 1992) and the receptor-like PTPase, LAR (leukocyte antigenrelated PTPase), catalyzed reaction (Cho et al, 1992). The ^^P NMR chemical shift of the phosphoenzyme intermediate is typical of a thiophosphate bond. Moreover, the formation and decay of the intermediate have been shown to be kinetically competent by fast quench technique

32

ZHONG-YIN ZHANG

(Cho et al., 1992). All of the above experiments point to the existence of a covalent cysteinylphosphate enzyme intermediate on the kinetic pathway. The involvement of a phosphocysteine intermediate in phosphatasecatalyzed hydrolysis reactions is novel. A thiophosphate linkage in proteins has been reported only in bacterial thioredoxin (Pigiet and Conley, 1978) and mannitol carrier-specific transporter enzyme II (Pas et al., 1991). The biological significance of thiophosphate in thioredoxin is not clear. A phosphocysteine intermediate proposed for the mannitolcarried specific transporter enzyme in bacteria may be required for phosphate transfer from phosphoenolpyruvate to mannitol. Thiophosphate esters have been suggested as "models" for intermediates in enzymatic phosphoryl transfers (Walsh, 1952), but evidence for their existence in enzymatic systems has been weak until now. Because the energy of the P-S bond (45-50 kcal/mol) is considerably less than that of the P - 0 bond (95-100 kcal/mol), P - S bond cleavage is much more facile than P - 0 cleavage (Bruice and Benkovic, 1966). Thus, it is not unreasonable for PTPases to utilize a thiophosphate as a covalent enzyme intermediate in catalysis. Previous work suggests that nonenzymatic thiophosphate ester hydrolysis exhibits many of the characteristics of its oxygen counterparts (Herr and Koshland, 1957; Dittmer et al, 1963; Bruice and Benkovic, 1966; Milstien and Fife, 1967). PTPases provide a unique system in which to examine the enzyme-catalyzed phosphoryl transfer where a cysteine nucleophile is directly involved. C. Invariant Arginine in PTPase Signature [\/lotif

As discussed above, the invariant Cys present in the PTPase signature motif, which includes the active site sequence (HyV)C(X)5R(S/T) (where X can be any amino acid; Fig. 4), is absolutely required for catalysis and is directly involved in phosphoenzyme formation. The importance of the invariant Arg residue in the PTPase signature motif (Arg-409 in Yersinia PTPase) has also been recognized. Mutations of the invariant Arg residue in the PTPase signature motif resulted in complete loss of enzymatic activity for two receptor-like PTPases, LAR and CD45 (Streuh et al, 1990; Johnson et al, 1992) and the low M, phosphatase from bovine liver (Cirri et al, 1993). However, from such studies, it is not clear whether the Arg residue is essential for PTPase folding and structure or catalysis, or both. The extraordinary reactivity of the Yersmia PTPase (Zhang et al, 1992) has made it possible to examine the effect of mutagenesis on the invariant Arg residue within the active site. The higher intrinsic activity of Yersinia PTPase has made it possible to measure ^cat and K^ values for both the R409A and


33

R409K site-directed mutants to address the functional significance of Arg-409 in terms of substrate binding and catalysis (Zhang et al., 1994d). A 8200-fold decrease in ^cat and a 26-fold increase in K^ were observed for the R409A mutant. Interestingly, the R409K mutant displayed a ^cat value identical to that of R409A, and the apparent K^ value for pNPP was only 1.9-fold higher than that of the wild-type enzyme. The R409A mutation decreases the arsenate binding affinity by 47-fold, while R409K decreases the arsenate binding affinity by 18fold. These results suggest that Arg-409 plays a critical role in phosphosubstrate binding. The most important function of Arg-409 may be transition state stabilization, which is reflected by the dramatic reduction in both ^cat/^m and ^cat values for the Arg-409 mutants. Tungstate is a competitive inhibitor of the Yersinia PTPase, with a binding constant of 61 IJLM at pH 7.0 and ionic strength of 0.15 M. The three-dimensional structure of the Yersinia PTPase-tungstate complex (Stuckey et al., 1994) shows that tungstate oxygen atom(s) ion pair with the positively charged guanidinium group of Arg-409 (Fig. 3). Tungstate oxygen atoms labeled a and b form hydrogen bonds with the Ns and NT/ of Arg-409. The tungstate oxygen atom denoted as c is hydrogen bonded to the NH amides of the peptide backbone making up the phosphate-binding loop. The tungstate oxygen atom denoted as d is projecting out of the active site pocket and most likely corresponds to the oxygen atom present in the scissile substrate phosphate ester bond. Figure 3 shows that the thiolate anion of Cys-403 is positioned at the base of the active site and would likely attack the phosphate ester, forming a trigonal bipyramidal transition state. The side chain of Arg-409 is in an ideal position for effective stabilization of the transition state (Fig. 3). Cleavage of the phosphoester linkage would then produce the Cys-403 thiophosphate intermediate with release of phenol/alcohol. Hydrolysis of the phosphoenzyme intermediate would most likely require water to attack in the same axial position from which the phenol/ alcohol departed. The attack of water on the phosphoenzyme intermediate would again form a trigonal bipyramidal transition which can be similarly stabilized by Arg-409. The subsequent cleavage of the thiophosphate bond would regenerate the thiolate anion of Cys-403 and yield the other product of the reaction, inorganic phosphate. Studies have shown that a guanidinium group (present in an arginine) is ideally suited for interaction with phosphate by virtue of its planar structure and its ability to form multiple hydrogen bonds with the phosphate moiety (Cotton et al., 1973). The ability of the guanidinium group to form a coplanar bidentate complex with two equatorial oxygen atoms present on the phosphate during catalysis provides a

34

ZHONG-YIN ZHANG

plausible mechanism for stabilization of the trigonal bipyramidal transition state(s). The geometry associated with the amino group of the alternate cationic Lys side chain would not be expected to be able to form a coplanar bidentate complex with the trigonal bipyramidal transition state, which may explain why the R409K mutant showed no improvement in ^cat when compared to R409A. The fact that a Lys residue at position 409 can partially replace the Arg residue in terms of substrate and inhibitor binding, while at the same time being unable to substitute for the Arg in catalysis, suggests that the transition state(s) likely employ the unique structural properties of the guanidinium side chain of Arg-409. It is most likely that the conserved Arg residue in the PTPase active site is geometrically positioned in such a manner that it interacts more favorably with the transition state than with the ground state. D. Conserved Serine/Threonine in PTPase Signature IViotif

In addition to the essential Cys and Arg residues, a conserved Ser or Thr can be found in the PTPase signature motif immediately after the invariant Arg residue (Fig. 4, Zhang et al., 1995c). The dual-specificity phosphatase Cdc25 is the only one that lacks a hydroxyl group at this position. Interestingly, Cdc25 is several orders of magnitude less reactive than other PTPases (Dumphy and Kumagai, 1991; Zhang et al, 1992). In the bovine low M, PTPase structure (Su et al, 1994; M. Zhang, et al., 1994), as well as in the Yersinia PTPase (Stuckey et al., 1994) and the human PTPIB (Barford et al, 1994) structures, the hydroxyl group of the conserved Ser/Thr is approximately 3 A to the Sy of the active site Cys residue (Fig. 5), making a reasonable good S-HO hydrogen bond (Gregoret et al, (1991). Because the PTPases, the dual-specificity phosphatases, and the low Mj. phosphatases effect catalysis through a covalent thiophosphate enzyme intermediate, the catalyzed reaction must be composed of at least two chemical steps, i.e., formation and breakdown of the phosphoenzyme intermediate. The phosphoryl group in the substrate is first transferred to the nucleophilic active site thiolate group of the enzyme to form the phosphoenzyme intermediate (^2)? which is then hydrolyzed by water (^3) (Scheme I). The kinetic scheme is composed of substrate binding, followed by two chemical steps. ki

E + ROPOf^

1 E • ROPOi

< E-P ROH

SCHEME 1

ko

- E + Pi


35

FIG. 5. Active site conformation of the PTPase signature motif corresponding to residues Cys-12 to Ser-19 in the bovine low M, PTPase (M. Zhang et al, 1994). The hydrogen bond between the sulfur atom of Cys-12 and the hydroxyl group of Ser-19 is highlighted. Reprinted with permission from Y. Zhao and Z.-Y. Zhang, Biochemistry 35,11797-11804 (1996). Copyright 1996 American Chemical Society.

phosphorylation {k^ and dephosphorylation (^3), where E is the enzyme, ROPO3 the substrate, E-ROPOi the enzyme-substrate Michaehs complex, E-P the phosphoenz3mae intermediate, ROH the phenol, and Pi inorganic phosphate. If the net rate of intermediate breakdown is slower than that of intermediate formation, one would predict a "burst" ofp-nitrophenol production using p-nitrophenyl phosphate (pNPP) as a substrate. Burst kinetics has been demonstrated with the Yersinia PTPase (Zhang eâZ., 1995c), rat PTPl (Zhang, 1995a), the dual-specificity phosphatase VHR (Zhang et al, 1995b), and the low M^ phosphatases (Zhang and Van Etten, 1991; Zhang et al, 1995a). This has permitted the determination of individual rate constants directly associated with the formation {k^ and breakdown (^3) of the phosphoenzyme intermediate. Thus, an evaluation of the burst kinetics combined with the technique of site-directed mutagenesis should allow one to ascertain specific contributions of active site residues to the individual steps of the phosphatasecatalyzed reaction. It was shown that under most conditions, the decomposition of the cysteinylphosphate enzyme intermediate (^3) is the rate-limiting step for the overall phosphatase-catalyzed hydrolysis. Supporting evidence includes that up to 27-74% of PTPase can be trapped as a covalent adduct using ^^P-labeled p-nitrophenyl phosphate (Zhang, 1990; Guan and Dixon, 1991; Wo et al, 1992) and that such an intermedi-

36

ZHONG-YIN ZHANG

ate can be prepared in sufficient amount for ^^P NMR analysis (Wo et al, 1992; Cho eâZ., 1992). Site-directed mutagenesis and pre-steady-state stopped-flow kinetic experiments have demonstrated that the conserved hydroxyl group in the PTPase signature motif plays a critical role in efficient E - P hydrolysis in the Yersinia PTPase (Zhang et al., 1995b), the dual-specificity phosphatase, VHR (Denu and Dixon, 1995), and the low Mr phosphatase-catalyzed reaction (Zhao and Zhang, 1996). It appears that the elimination of the hydroxyl group in the conserved Ser/Thr has only a modest effect on k^, whereas its major impact seems to be primarily reflected in ^3. For example, in the reaction catalyzed by the low M^ phosphatase (Stpl) from the fission yeast S. pombe (Mondesert et al, 1994), elimination of the hydroxyl group at Ser-18 decreases the rate of E - P formation (^2) and breakdown (ks) by 4.3- and 35.7-fold, respectively (Zhao and Zhang, 1996). Similarly, substitution of the corresponding Thr residue by an Ala in the Yersinia PTPase resulted in a 2.4- and 30.4-fold reduction in ^2 and k^, respectively, at pH 5.8 (Zhang et al., 1995c). Figure 6 shows the differential effects of Ala substitution at residue Thr-410 of the Yersinia PTPase on ^2 and kz at pH 6.0 (Zhang et al, 1995c). Thus, results from the Yersinia PTPase, the dual-specificity 0.25

E

c o ^

0.2

0.15

0

o c

CO

0.1

o

se-

An

1

1 B ** 1 1 ^3 fS

LU

X

1 11

-7 -6 -7 -6 E-64a CA-074

VD3

T 1

II

^

1

B ^11 1

III i 1 ^^ 1 11 " i f--

-7 -6 -7 -6 E-64a CA-074

ITNF-a

fl

1

^H igi^ I j

O S

Jl

d1 [ 1

-7 -6 -7 -6 E-64a

CA-074

iPTHi

FIG. 6. Induction of pit formation by addition of PTH, la,25-(OH2D3, and TNF-o; and the suppression by E-64a or CA-074. All values are given as the percent of control with PTH alone. The numbers - 7 and - 6 represent an inhibitor concentration of lO""^ and 10"^. * p < .05, ** p < .01: significant differences from control group.

V. Cathepsin L Secreted from Osteoclasts as Precursor Form and Processed by Cysteine Proteinase(s) in Bone Lacunae The 39-kDa cysteine proteinase, which seems to be a precursor form, was secreted into culture medium of a bone cell mixture during the process of pit formation. Procathepsin L purified from rat long bones under cold alkaline conditions was rapidly converted to the mature form under acidic conditions at room temperature. However, this conversion was inhibited by the addition of E-64 during the process, suggesting that the procathepsin L secreted into lacunae (10) is autocatalytically converted to the mature form, as shown in the Western blot in Fig. 7. All procathepsins can be activated by the same processes.

188

NOBUHIKO KATUNUMA

kDa

49.5

•

-

;,-•

,

."

;xi

-

•

" I

I

;52. 5

27.5 pK

A

B

FIG. 7. Inhibition of processing of procathepsin L by addition of E-64 during purification and detection using Western blotting analysis by anti-cathepsin L antibody. Procathepsin L was processed in 0.1 M acetate buffer, pH 4.5, for 30 min in the presence of 10"^ M E-64 (lane A) and the absence (lane B) of E-64.

VI. Inhibitory Mechanisms of H+-ATPase, Carbonic Anhydrase II, and Monensin on Pit-Forming Assay Bone resorption depends on the secretion of proton and procathepsin L from osteoclasts into the extracellular lacunae. The proton is generated by carbonic anhydrase II in osteoclasts and activity in the lacunae by a proton pump driven by vacuolar-type H+-ATPase at the ruffled border, and the secreted proton participates in the bone resorption. However, the interrelated regulation between the proton and procathepsin L secretions from osteoclasts is still unclear. Addition of Bafilomycin Ai, an H+-ATPase inhibitor, completely suppressed pit formation stimulated by PTH but did not suppress procathepsin L secretion.

189

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monensin FIG. 8. Influences of monensin on bone resorption number induced by PTH and Z-PheArg-MCA hydrolytic activities of rat osteoclasts. Concentrations of PTH and monesin was 5 X 10"^ M. Each value of pit formation indicates the mean ± SD offiveobservations. **p < .01; significant difference from monensin minus group (Student's ^test).

Administration of acetazolamide, a carbonic anhydrase II inhibitor, suppressed both bone resorption and procathepsin secretion. These findings suggest that the secretion of procathepsin L and the activation (processing) of the secreted procathepsin L may be regulated by production and secretion of proton in the osteoclasts (12). The effects of monensin on bone resorption induced by PTH and ZPhe-Arg-MCA hydroljîc activity in isolated osteoclasts were assayed. Figure 8 shows the inhibition of pit formation and proteolytic activity in osteoclasts (10). Monensin itself does not inhibit cysteine proteinase activity, but it does inhibit the targeting of cysteine proteinases into lysosomes to prevent the excretion of cathepsins. These findings confirm that lysosomal cysteine proteinase in osteoclasts secreted into the lacunae plays an important role in the process of bone resorption.

VII. Suppression of Bone Resorption by Cathepsin L Family Inhibitors in Vivo The results of pit formation in vitro reflex directly affect the serum calcium level derived from bone resorption in vivo. We examined the in vivo effects of E-64, cystatin A, and CA-074 on the changes in serum

190

NOBUHIKO KATUNUMA

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Time (hr) FIG. 9. Effect of E-64 or CA-074 on serum calcium levels of rats on low-calcium diets. (•) No inhibitor; (A) 3 mg/100 g body weight of inhibitor; (•) 6 mg/100 g body weight of inhibitor. Levels measured 0-10 hr after inhibitor administration. Each value indicates the mean ± SEM of five observations, p < .05; significant difference from control (Student's ^test).

calcium of rats on a low-calcium diet and also with experimental malignant hypercalcemia as shown in Figs. 9 and 10, respectively. At 3 and 6 hr after the injection of E-64 (doses of 3 and 6 rag/ 100 g body weight) into rats on a low-calcium diet, serum calcium was significantly decreased in a dose-dependent manner. Administration of cystatin A (dose of 8 mg/100 g body weight) also significantly decreased the calcium level in the serum. At 1 hr after injection of E-64 or cystatin A, all cysteine proteinases in femur bone were inhibited effectively. By contrast, CA-074 did not affect the serum calcium level, although cathepsin B activity in the bone was specifically inhibited. These findings reconfirmed in vivo that cathepsin L rather than cathepsin B plays a central role in bone collagen degradation.

Vlli. Possible Strategies for New Drug Design to Protect Bone Resorption I propose the following two kinds of strategy to protect bone resorption. For suppression of cathepsin L secretion from osteoclasts: (1) inhibit the PTH effect mediated by the receptor located on the membrane of osteoblasts, (2) inhibit the second messenger information

191

BONE RESORPTION BY OSTEOCLASTS

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from osteoblast to osteoclast, (3) suppress procathepsin L secretion by administering a monensin-like drug, and (4) suppress proton formation and secretion by inhibition of the ATP-proton pump and of carbonic anhydrase. For inhibition of cathepsin L activity: (1) inhibit cathepsin L activity by a specific inhibitor and (2) inhibit conversion from precursor to the mature form. ACKNOWLEDGMENT I sincerely t h a n k Miss Yoshiko Nagai for preparation of this manuscript.

REFERENCES 1. Delaisse, J. M., Boyde, A., Maconnachie, E., Ali, N. N., Sear, C. H. J., Eedkhout, Y., Vaes, G., and Jones, S. J. (1987). Bone 8, 305.

192

NOBUHIKO KATUNUMA

2. Delaisse, J. M., and Vaes, G. (1992). In "Biology and Physiology of the Osteoclast" (B. R. Rifkin and C. V. Gay, eds.), p. 290. CRC Press, Boca Raton, FL. 5. Towatri, T., Hanada, K., Katunuma, N., et al. (1991). FEES Lett. 280, 311. 4. Katunuma, N., and Kominami, E. (1995). In "Methods in Enzymology" Vol. 251, Chaper 37, pp. 382-397. Academic Press, San Diego, CA. 5. Lenarcic, B., Ritonja, A., Dotenc, J., Stoka, V., Berbic, S., Pungercar, J., Stukelj, B., and Turk, V. (1993). FEES Lett. 336, 289. 6. Kakegawa H., Nikawa, T., Tagami, K, Kamioka, T., Drobnic-Kosorok, M., Lenarcic, B., Turk, v., and Katunuma, N. (1993). FEES Lett. 321, 247. 7. Katunuma, N., Kakegawa, H., Matsunaga, Y., Nikawa, T., and Kominami, E. (1993). Agents Actions, Suppl. 42, 195. 8. Turk, D., Podobnik, M., Popovic, T., Katunuma, N., Bode, W., Huber, R., and Turk, V. (1995). Biochemistry 34(14), 4791-4797. 9. Tagami, K, Kakegawa, H., Kamioka, H., Sumitani, K., Kawata, T., Lenarcic, B., Turk, v., and Katunuma, N. (1994). FEES Lett. 342, 308. 10. Kakegawa, H., Tagami, K, Ohba, Y., Sumitani, K., Kawata, T., and Katunuma, N. (1995). FEES Lett. 370, 78-82. 11. Inubushi, T., Kakegawa, H., Kishino, Y., and Katunuma, N. (1994). J. Eiochem. (Tokyo) 166, 282-284. 12. Ohba, Y., Yamamoto, T., and Katunuma, N. (1996). FEES Lett. 387, 175-178.

Indx

Abdominal muscles, apoptosis in, 81 Acetazolamide, effect on bone resorption, 189 Acid phosphatases, cataljîc mechanism of, 31 Aconitase, iron-sulfur cluster of, 14 Acquired immunodeficiency syndrome. See AIDS Actinomycin D, apoptosis induction by, 70 Adenosine triphosphate (ATP), biochemical importance of, 149 Aging apoptosis role in, 70, 77, 107-121 benefits of caloric restriction in, 114-115 elevated levels of reactive oxygen species in, 123, 143 immune function decline in, 112-114 osteoporosis as problem of, 179 Agrobacterium rhizogenes, rolB oncogene from, 24 AIDS apoptosis in, 107 Fas-mediated killing in, 89 N F - K B role in, 150, 153-156, 157 Alkaline phosphatases, catalytic mechanism of, 31 Alzheimer's disease (AD), apoptosis and, 107, 115, 117 Amyotrophic lateral sclerosis (ALS), apoptosis and, 107, 115, 117 Antibiotics, bacterial resistance to, 174, 175 Antioxidants, in organisms, 149 Antipain, as cysteine proteinase inhibitor, 179 AP-1, redox sensitivity of, 131-132, 141 APO-1. See Fas and Fas ligand Apopain. See Yama Apoptosis accelerants for, 72, 76 aging and, 70, 77, 107-121 antagonists of, 72, 75, 76, 80 caloric restriction and, 114-115 cell response in, 71-72

cell senescence and, 110-112 description of, 69 Fas-mediated, 24, 69-105, 110, 112-114 function of, 69-70, 107-108 genes involved in, 108-110 immunological aging and, 112-114 induction of, 7 0 - 7 1 neurodegenerative disease and, 115-116 nucleosomal ladder of, 69, 70, 71, 80, 81, 91 proteolysis in, 83 regulation of, 7 2 - 9 1 Apoptotic bodies, formation of, 69, 71 Aprotinin, as serine proteinase inhibitor, 183 Arachidonic acid, oxidative damage to, 123 Arginine oxidative damage to, 124 in PTPase signature motif, 22, 27, 28, 30,32-34,38,46 Articular chondrocytes, apoptosis in, 117 Ascorbate, as free radical scavenger, 123 Aspartic acid in ICE-cleaved substrates, 84 in PTPases, 37, 38, 39, 40, 46-47, 61 ATF/CREB family, DNA binding of, 141, 142 Autoimmune diseases apoptosis and, 107 defective signaling network in, 21 increase in aging, 113 PTPase role in, 21, 57 B Bacillus suhtilis, oxidative stress responses of, 164, 173-174 Bacteria antibiotic resistance of, 174, 175 oxidative stress responses of, 163-177 Bad, as accelerant for apoptosis, 76 BAG-1, as antagonist of apoptosis, 76 Bak, as accelerant for apoptosis, 76 Bax, as accelerant of apoptosis, 73-75 193

194 Bcl-2 as antagonist of apoptosis, 7 2 - 7 3 , 91, 123 Mcl-l interaction with, 76 Bcl-2 family control of expression of, 77-78 list of members of, 72 as regulators of apoptosis, 72-78, 92, 111 bcl-2 gene, role in apoptosis, 109, 111, 113, 116, 117 Bcl-x, as antagonist of apoptosis, 75 Benzylphosphonic acid, as PTPase inhibitor, 59 Bilirubin, as oxygen scavenger, 134-135 B lymphocytes, apoptosis in, 118 Bone resorption drug protection of, 190-191 mechanism of, 181 pit formation test for, 180, 181, 182, 183, 187 procathepsin L regulation of, 179-191 suppression by cathepsin L inhibitors, 180-183 Brain, apoptosis in, 116-117 Breast carcinoma, apoptosis in, 87 Burst kinetics, of phosphatase activity, 35, 36 BZLFl transcription factor, redox sensitivity of, 133

CA-074, effect on bone resorption, 189-190 Caenorhadbitis elegans, apoptosis studies on, 70, 76, 108, 109, 116 Calcitonin, suppression of cathepsin release by, 180 Caloric restriction (CR), in upregulation of apoptosis, 114-115, 118 Cancer apoptosis role in control of, 107, 117-118 defective signaling network in, 21 deregulated protein-tyrosine kinase role in, 23 elevated levels of reactive oxygen species in, 123 metastatic, NF-zcB role in, 150, 1516-157 PTPase role in, 21, 57

INDEX Carbonic anhydrase II, effect on bone resorption, 188 ^-Carotene, as free radical scavenger, 123 Catalase, antioxidant reactions catalyzed by, 123 Cataracts, IRE mutation role in, 13 Cathepsins, inhibitors of, 179, 180 Cathepsin L regulation of secretion of, 184-187 suppression of bone resorption by inhibitors of, 180-183, 189-190 c-cdc gene, role in apoptosis. 111 CD45 biological function of, 24 as receptor-like PTPase, 22, 32 Cdc25, as dual-specificity phosphatase, 24 CD95 protein. See Fas and Fas ligand Ced-3, role in apoptosis, 78-79 ced genes, role in apoptosis, 76, 108, 113 Cell death. See also Apoptosis necrosis as, 107 Cell growth and differentiation PTPase role in, 21, 62 redox sensitivity of, 138 Cell senescence, apoptosis and, 110-112 Cell signaling, antioxidant role in, 149 Ceramide, role in apoptosis, 84-86, 91, 92, 110 c-fos/c-jun gene, role in signal transduction, 131-132 c-fos gene expression of, 137 role in apoptosis. 111 Chemotherapeutic agents, apoptosis induction by, 70, 107 Chromosome 11, H chain gene on, 2 Chromosome 19, L chain gene on, 2 Chromosomes 14 and 18, bcl-2 gene and, 109 Cinnamic acid, structure of, 60 c-Jun transcription factor, activation of, 137 c-myc gene, role in apoptosis, 110, 111 Colonic epithelium, apoptosis in, 74 corkscrew gene, 24 Corticosteroids, apoptosis induction by, 70 Covalent phosphoenzyme intermediate, in PTPase kinetic pathway, 63

195

INDEX Cowpox virus, CrmA of, 82 CrmA, as ICE inhibitor, 82 Cubanes, iron-sulfur clusters as, 4, 14 Cu/Zn-superoxide dismutase, deleterious mutation affecting, 117 Cyclins, role in cell division, 92 Cycloheximide, apoptosis induction by, 70,73 Cystatin, as cathepsin inhibitor, 181, 189 Cysteine in degradation domain of iron regulatory protein 2, 19 in PTPase signature motif, 22, 2 7 - 2 8 , 30, 31, 38, 46 Cysteine proteinase inhibitors, bone-resorption inhibition by, 179 Cysteinylphosphate enz5mrie, in phosphatase pathway, 32 Cytokines apoptosis induction by, 70 N F - K B in regulation of, 150, 151, 157

Cjrtomegaloviruses, N F - K B regulation of, 150 C5ôplasm, apoptosis effects on, 69 D Death domain, of Fas, 87 Death proteases definition of, 82 role in apoptosis regulation, 78, 79, 8 0 - 8 1 , 88, 92 substrates for, 8 3 - 8 4 Death signaling cascade, propagation of, 81, 82, 86 Degenerative diseases, elevated levels of reactive oxygen species in, 123 Diabetes defective signaling network in, 21 PTPase role in, 21, 57 Diacylglycerol, as ceramide inhibitor, 8 4 - 8 5 , 86 Disease apoptosis in, 73, 87, 89 defective signaling network in, 21 elevated levels of reactive oxygen species in, 123 IRE mutation role in, 13 DNA apoptosis effects on, 69, 70, 110

oxidative damage to, 1, 115, 124, 125, 143, 163, 164 D128 mutant, acid-base catalysis by, 40 D129 mutant, acid-base catalysis by, 39 Double-displacement pathway, of phosphatase action, 30 Drosophila apoptosis in, 110 succinate hydrogenase of, 15, 16 Dual-specificity phosphatases, catalytic mechanism of, 2 8 - 2 9 E eALAS protein, IRE-IRP regulation of, 14, 16 E g r l transcription factor activation of, 137 redox sensitivity of, 129 E - P formation and E - P breakdown steps, in PTPase-catalyzed reactions, 42-46 Epstein-Barr virus protein, as apoptosis inhibitor, 77 Erythrocyte amino levulinic acid synthase, role in heme biosynthesis, 4 Escherichia coli aconitase of, 8, 9, 14 oxidative stress response in, 125, 163, 164-169, 172, 174 E-selectin, N F - K B regulation of, 150 E-64, as cysteine proteinase inhibitor, 179, 180, 181, 189 Eukaryotes, iron-metabolism regulation in, 1-19 Evolution, iron-sulfur cluster role in, 13-14 F FADD, reaction with Fas death domain, 88 Fas and Fas ligand apoptosis mediated by, 24, 69-105, 110, 112-114 expression of, 88-89 structure and activity of, 87-89 Fas-associated phosphatase (FAP-1), biological function of, 24 fas gene, overexpression of, 113 Fenton reaction, 1, 135

196 Ferritin expression of, 5 role in oxidant stress reversal, 135 structure of, 3 translational regulation of, 1-3, 4, 16 Ferritin repressor protein, 3. See Iron regulatory protein 1 (IRPl); Iron regulatory protein 2 (IRP2) Ferrous iron, in Fenton reaction, 1 Fibroblasts apoptosis in, 111-112, 117, 118 senescent, in aged persons, 114 FLICE, role in apoptosis, 82 Flucocorticoids, role in ced activation, 109-110 4-( Fluoromethyl)phenylphosphate (FMPP), as PTPase inhibitor, 6 0 - 6 1 fra genes, expression of, 137 Free radicals. See also Reactive oxygen species (ROS) neuronal death from, 108 G gadd45 gene, transcription of, 137 Gene transcription, PTPase role in, 21 Genome, oxidant radical effects on, 137 gld gene, role in apoptosis, 113 Gliomas, apoptosis in, 91 Glucocorticoid receptor (GR), redox sensitivity of, 127 Glutamate-ammonia ligase, degradation of, 10 Glutamic acid, in PTPases, 37, 38, 40, 51 Glutamine synthetase of E. coli, degradation of, 10 as target of reactive oxygen, 115 Glutaredoxin, in cell signaling, 149 Glutathione, as free radical scavenger, 123 Glutathione peroxidase, antioxidant reactions catalyzed by, 123 Glycerophosphate, as PTPase substrate, 56 Glycine, in degradation domain of iron regulatory protein 2, 19 Gold ion, NF-/cB blockage by, 154 G proteins, in signal transduction, 92 Granzymes, apoptosis induction by, 70, 81 Growth factors effect on apoptosis, 7 0 - 7 1 , 111, 117 role in ced activation, 109-110

INDEX Guanidium group, in arginine of PTPase signature motif, 3 3 - 3 4 H Haemophilus influenzae, oxidative stress responses of, 164, 171 H+-ATP, effect on bone resorption, 188 H chain, of ferritin, 2 Heat shock, oxidant stress and, 136 Heat shock factor (HSF), oxidation role in activation of, 131 Hematopoietic tissue, Bcl-x expression in, 75 Heme biosynthesis, eALAS protein role in, 14 Hinge-linker, in mitochondrial aconitase binding site, 7, 14 Hippocampus, apoptosis in, 116-117 Histidine, oxidative damage to, 124 HIV. See AIDS; Human immunodeficiency virus (HIV) HIV-1 Tat, upregulation of Fas by, 89 Homeoboxes, embryogenesis and, 92 HoxB5, redox sensitivity of, 132 H u m a n immunodeficiency virus (HIV) anti-HIV drug screening for, 158 NF-/cB role in, 136, 150, 154-156, 157 Hydrogen peroxide cell damage from, 115, 135, 137 ferrous iron reaction with, 1 Hydroperoxidases, of bacteria, 164, 171 Hydroperoxyl radical, as reactive oxygen species, 123, 163 8-Hydroxy-2'-deoxyguanosine, as marker for DNA oxidation, 124 Hydroxyl radical cell damage from, 115 formation in Fenton reaction, 1 as reactive oxygen species, 123, 163 Hyperferritinemia-cataract syndrome, IRE mutation role in, 13

ICAM-1, NF-/

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