Colloquium on Proteolytic Processing and Physiological Regulation

Table of contents :
COLLOQUIUM ON PROTEOLYTIC PROCESSING AND PHYSIOLOGICAL REGULATION
NATIONAL ACADEMY OF SCIENCES
Proteolytic Processing and Physiological Regulation
A COLLOQUIUM SPONSORED BY THE NATIONAL ACADEMY OF SCIENCES
FEBRUARY 20—21, 1999
Saturday, February 20, 1999
Sunday, February 21, 1999
PROCEEDINGS OF THE NATIONAL ACADEMY OF SCIENCES OF THE UNITED STATES OF AMERICA
Contents
National Academy of Sciences Colloquia
BOUND REPRINTS AVAILABLE
Proteolytic enzymes, past and future
Caspase activation: The induced-proximity model
Structural aspects of activation pathways of aspartic protease zymogens and viral 3C protease precursors
Conversion of Gastric Aspartic Protease Zymogens
Conversion of Proplasmepsin II
Autocatalytic Excision of Picornaviral 3C Proteases
The catalytic sites of 20S proteasomes and their role in subunit maturation: A mutational and crystallographic study
MATERIALS AND METHODS
RESULTS AND DISCUSSION
The structure of the human βII-tryptase tetramer: Fo(u)r better or worse
CONCLUSION
Sonic hedgehog protein signals not as a hydrolytic enzyme but as an apparent ligand for Patched
MATERIALS AND METHODS
RESULTS
DISCUSSION
Structure-assisted design of mechanism-based irreversible inhibitors of human rhinovirus 3C protease with potent antiviral...
Picornaviral 3C Proteases
Inhibitors of 3C Protease and the Issue of Serotypic Diversity Among Rhinoviruses
Irreversible Michael Acceptors as Inhibitors of 3C Protease
Michael-Acceptor Inhibitors of 3C Protease: Structure-Activity Studies
AG7088, a 3C Protease Inhibitor with Potent Antiviral Activity Against Multiple Human Rhinovirus Serotypes
Kinetic stability as a mechanism for protease longevity
Cysteine protease inhibitors as chemotherapy: Lessons from a parasite target
METHODS
RESULTS
DISCUSSION
How the protease thrombin talks to cells
How Does a Protease Talk to a Cell?
Irreversible Activation, Disposable Receptors, and Intracellular Reserves
A Protease-Activated Receptor Family
PARs and Platelet Activation
A Role for Thrombin Signaling in Embryonic Development and Other Processes?
Summary
VanX, a bacterial D-alanyl-D-alanine dipeptidase: Resistance, immunity, or survival function?
Chaperone rings in protein folding and degradation
Architecture-Function Considerations
Substrate Protein Recognition
Action of ATP
Commitment of Substrate
Prospects for Further Mechanistic Understanding
A proteolytic pathway that controls the cholesterol content of membranes, cells, and blood
Two-Step Proteolytic Release of SREBPs
SREBP Cleavage-Activating Protein (SCAP)
SCAP as a Sterol Sensor
Candidate Gene for Site-2 Protease
Candidate Gene for Site-1 Protease
Unresolved Questions
Cellular mechanisms of β-amyloid production and secretion
Reverse biochemistry: Use of macromolecular protease inhibitors to dissect complex biological processes and identify a...
MATERIALS AND METHODS
RESULTS
DISCUSSION

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COLLOQUIUM ON PROTEOLYTIC PROCESSING AND PHYSIOLOGICAL REGULATION

NATIONAL ACADEMY OF SCIENCES WASHINGTON, D.C. 1999

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NATIONAL ACADEMY OF SCIENCES

Colloquium Series In 1991, the National Academy of Sciences inaugurated a series of scientific colloquia, five or six of which are scheduled each year under the guidance of the NAS Council’s Committee on Scientific Programs. Each colloquium addresses a scientific topic of broad and topical interest, cutting across two or more of the traditional disciplines. Typically two days long, colloquia are international in scope and bring together leading scientists in the field. Papers from colloquia are published in the Proceedings of the National Academy of Sciences (PNAS).

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PROTEOLYTIC PROCESSING AND PHYSIOLOGICAL REGULATION

Proteolytic Processing and Physiological Regulation

A COLLOQUIUM SPONSORED BY THE NATIONAL ACADEMY OF SCIENCES FEBRUARY 20–21, 1999 Saturday, February 20, 1999 Hans Neurath, University of Washington Welcome and introduction: Proteolytic enzymes, past and future David Agard, University of California, San Francisco Kinetic stability and folding of proteases: twin paradigms for protease pro regions Michael James, University of Alberta Structural basis and mechanism of zymogen activation David Matthews, Agouron Pharmaceuticals, Inc. Structure-assisted design of mechanism based irreversible inhibitors of human rhinovirus 3C protease with potent antiviral activity against multiple rhinovirus serotypes Christopher Walsh, Harvard University Role of D, D-Peptidase in Vancomycin Resistance Earl Davie, University of Washington Introduction to Protease activated receptors Shaun Coughlin, University of California, San Francisco Thrombin signaling: Molecular mechanisms and roles in vivo Vishva Dixit, Genentech, Inc. Identification of components of the cell death pathway Wolfram Bode, Max-Planck-Institute for Biochemistry Structure of tryptase, a cage-like serine proteinase involved in asthma, allergic and inflammatory disorders Philip Beachy, Johns Hopkins University Hedgehog protein biogenesis and signaling Marc Kirschner, Harvard University The role of proteases in the regulation of cell cycle

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PROTEOLYTIC PROCESSING AND PHYSIOLOGICAL REGULATION

Sunday, February 21, 1999 C.S.Craik, University of California, San Francisco Introduction Arthur Horwich, Yale University Chaperone Rings in Protein Folding and Degradation Robert Huber, Max-Planck-Institute for Biochemistry Structure of the archaeal and yeast 20S proteasomes and of the eubacterial Analog HslV Sukanto Sinha, Athena Neurosciences Cellular mechanism of beta amyloid production and secretion Michael Brown, University of Texas Southwestern Medical Center A proteolytic system that controls cholesterol metabolism Michael Brown Introduction Charles Craik, University of California, San Francisco Reverse biochemistry-using protease inhibitors to dissect complex biochemical processes Christine Debouck, Smith-Kline and Beecham Pharmaceuticals From genomics to drugs—cathepsin K and osteoporosis James McKerrow, University of California, San Francisco Parasite proteases—windows on molecular evolution and targets for drug design Joshua Boger, Vertex Pharmaceuticals Recognizing a drug

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TABLE OF CONTENTS

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PROCEEDINGS OF THE NATIONAL ACADEMY OF SCIENCES OF THE UNITED STATES OF AMERICA

Table of Contents

Papers from a National Academy of Sciences Colloquium on Proteolytic Processing and Physiological Regulation

Proteolytic enzymes, past and future Hans Neurath

10962–10963

Caspase activation: The induced-proximity model Guy S.Salvesen and Vishva M.Dixit

10964–10967

Structural aspects of activation pathways of aspartic protease zymogens and viral 3C protease precursors Amir R.Khan, Nina Khazanovich-Bernstein, Ernst M.Bergmann, and Michael N.G.James

10968–10975

The catalytic sites of 20S proteasomes and their role in subunit maturation: A mutational and crystallographic study Michael Groll, Wolfgang Heinemeyer, Sibylle Jäger, Tobias Ullrich, Matthias Bochtler, Dieter H.Wolf, and Robert Huber

10976–10983

The structure of the human βII-tryptase tetramer: Fo(u)r better or worse Christian P.Sommerhoff, Wolfram Bode, Pedro J.B.Pereira, Milton T.Stubbs, Jörg Stürzebecher, Gerd P.Piechottka, Gabriele Matschiner, and Andreas Bergner

10984–10991

Sonic hedgehog protein signals not as a hydrolytic enzyme but as an apparent ligand for Patched Naoyuki Fuse, Tapan Maiti, Baolin Wang, Jeffery A.Porter, Traci M.Tanaka Hall, Daniel J.Leahy, and Philip A.Beachy

10992–10999

Structure-assisted design of mechanism-based irreversible inhibitors of human rhinovirus 3C protease with potent antiviral activity against multiple rhinovirus serotypes D.A.Matthews, P.S.Dragovich, S.E.Webber, S.A.Fuhrman, A.K.Patick, L.S.Zalman, T.F.Hendrickson, R.A.Love, T.J.Prins, J.T.Marakovits, R.Zhou, J.Tikhe, C.E.Ford, J.W.Meador, R.A.Ferre, E.L.Brown, S.L.Binford, M.A.Brothers, D.M.DeLisle, and S.T.Worland

11000–11007

Kinetic stability as a mechanism for protease longevity Erin L.Cunningham, Sheila S.Jaswal, Julie L.Sohl, and David A.Agard

11008–11014

Cysteine protease inhibitors as chemotherapy: Lessons from a parasite target Paul M.Selzer, Sabine Pingel, Ivy Hsieh, Bernhard Ugele, Victor J.Chan, Juan C.Engel, Matthew Bogyo, David G.Russell, Judy A.Sakanari, and James H.McKerrow

11015–11022

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TABLE OF CONTENTS

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How the protease thrombin talks to cells Shaun R.Coughlin

11023–11027

VanX, a bacterial D-alanyl-D-alanine dipeptidase: Resistance, immunity, or survival function? Ivan A.D.Lessard and Christopher T.Walsh

11028–11032

Chaperone rings in protein folding and degradation Arthur L.Horwich, Eilika U.Weber-Ban, and Daniel Finley

11033–11040

A proteolytic pathway that controls the cholesterol content of membranes, cells, and blood Michael S.Brown and Joseph L.Goldstein

11041–11048

Cellular mechanisms of β-amyloid production and secretion Sukanto Sinha and Ivan Lieberburg

11049–11053

Reverse biochemistry: Use of macromolecular protease inhibitors to dissect complex biological processes and identify a membrane-type serine protease in epithelial cancer and normal tissue Toshihiko Takeuchi, Marc A.Shuman, and Charles S.Craik

11054–11061

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NATIONAL ACADEMY OF SCIENCES COLLOQUIA

vii

National Academy of Sciences Colloquia

BOUND REPRINTS AVAILABLE In 1991, the National Academy of Sciences (NAS) inaugurated a series of scientific colloquia, several of which are held each year under the auspices of the NAS Coun cil Committee on Scientific Programs. These colloquia address scientific topics of broad and topical interest that cut across two or more traditional disciplines. Typically two days long, these colloquia are international in scope and bring together leading scientists in the field. Papers presented at these colloquia are published in the Proceedings of the National Academy of Sciences (PNAS) and are available online (www.pnas.org). Because they have generated much interest, these papers are now available in the form of collected bound reprints, which may be ordered through the National Academy Press. Currently available are: Carbon Dioxide and Climate Change ($11) Held November 13–15, 1995 (Irvine, CA) Computational Biomolecular Science ($16) Held September 12–13, 1997 (Irvine, CA) Earthquake Prediction ($16) Held February 10–11, 1995 (Irvine, CA) Elliptic Curves and Modular Forms ($7) Held March 15–17, 1996 (Washington, DC) Genetic Engineering of Viruses and Viral Vectors ($21) Held June 9–11, 1996 (Irvine, CA) Genetics and the Origin of Species ($8) Held January 31-February 1, 1997 (Irvine, CA) Geology, Mineralogy, and Human Welfare ($11) Held November 8–9, 1998 (Irvine, CA) Neurobiology of Pain ($8) Held December 11–13, 1998 (Irvine, CA) Neuroimaging of Human Brain Function ($17) Held May 29–31, 1997 (Irvine, CA) Plants and Population: Is There Time? ($8) Held December 5–6, 1998 (Irvine, CA) Protecting Our Food Supply: The Value of Plant Genome Initiatives ($13) Held May 29–31, 1997 (Irvine, CA) Science, Technology, and the Economy ($12) Held November 20–22, 1995 (Irvine, CA) The Age of the Universe, Dark Matter, and Structure Formation ($13) Held March 21–23, 1997 (Irvine, CA)

Papers from future colloquia will be available for purchase after they appear in PNAS. Shipping and Handling Charges: In the U.S. and Canada please add $4.50 for the first reprint ordered and $0.95 for each additional reprint. Ordering Information: Telephone orders will be accepted only when charged to VISA, MasterCard, or American Express accounts. To order, call toll-free 1–800–624–6242 or order online at www.nap.edu and receive a 20% discount.

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PROTEOLYTIC ENZYMES, PAST AND FUTURE

10962

Proteolytic enzymes, past and future

This paper is the introduction to the following papers, which were presented at the National Academy of Sciences colloquium “Proteolytic Processing and Physiological Regulation,” held February 20–21, 1999, at the Arnold and Mabel Beckman Center in Irvine, CA. HANS NEURATH* Department of Biochemistry, Box 357350, University of Washington, Seattle, WA 98195 ABSTRACT Today’s knowledge is based on yesterday’s research, which, for me, started some 60 years ago. In the introduction to this colloquium, the past history of proteolytic enzymes is briefly reviewed against the background of simultaneously developing concepts and methodologies in protein chemistry. This history is followed by a sketch of more recent developments of the role of proteolytic enzymes in physiological regulation and an outlook of future trends apparent from current research. The history of proteolytic enzymes is intimately interwoven with that of protein chemistry. In the very early days, proteolytic enzymes were considered an impediment that had to be removed in the isolation of proteins generally. When I entered the field some 60 years ago, Northrop, Kunitz, and Herriott (1) had published the first edition of their treatise Crystalline Enzymes and demonstrated that, contrary to some prevailing notions, the crystalline proteolytic enzymes and protease inhibitors that they had isolated were chemical entities of constant solubility and hence obeyed the thermodynamic criteria of pure compounds. These compounds included pepsinogen, pepsin, and pepsin inhibitor, chymotrypsin, trypsin, their zymogens and inhibitors, carboxypeptidase, ribonuclease, hexokinase, diphtheria antitoxin, and a few others. Because these proteins were commercially unavailable, anyone interested in studying them had to isolate them the hard way. The field lay relatively dormant and awaited the development of more effective and specific methods of isolation, purification, and characterization of proteins, which came some 20 years later, including the methods of chromatography, gel electrophoresis, gel filtration, ultracentrifugation, amino acid analysis, and protein sequencing (2). In an effort to avoid the complexity of protein substrates, low molecular-weight synthetic peptides and their ester analogs were synthesized and found to simulate the specificity requirements of these proteases. Other landmarks included the discovery of natural and synthetic protease inhibitors such as disopropylfluoro phosphate, which introduced an organic phosphate label into the active site of serine proteases. Chemical characterization of active sites together with x-ray structure analysis of proteases showed that they can be grouped into families of common mechanism, similar structural features, and hence common evolutionary origin. They included the well known families of serine, cysteine, aspartic, and metallo endo- and exopeptidases. The number of proteases under investigation in the early days is minuscule compared with the current inventory of several thousand proteolytic enzymes that are coded by 2% of the structural gene pool (3). Interest in proteases was considerably stimulated by the recognition that, aside from their digestive action, proteases are involved in the regulation of a great many physiological processes. In many cases, regulation is mediated by the association of proteases with nonproteolytic domains that confer specificity to their interaction with receptor sites. The most studied among them are the proteases involved in blood coagulation, fibrinolysis, the complement system, and the processing of protein hormone precursors by specific convertases. A telling case of such an association is enterokinase, a protease that fulfills the simple but specific task of cleaving the amino-terminal hexapeptide during the activation of trypsinogen. Although enterokinase was discovered more than 50 years ago, it was only recently that its x-ray structure was elucidated by cloning and expressing the heavy chain (4). Surprisingly, it was found to be composed of a trypsin-like catalytic domain covalently bound to a series of nonprotease domains that also exist in unrelated proteins. One of these resembles the low density lipoprotein receptor, another resembles meprin, a third occurs in complement C1r, and yet another occurs in a macrophage receptor. The functional significance of these specific combinations is unknown. The term “limited proteolysis” was coined by Linderstrom-Lang to differentiate the restricted specificity of certain enzymes under certain conditions from the random proteolysis accompanying protein degradation. Proteolytic processing can be limited by the specificity of the protease, the accessibility of the susceptible peptide bond of the substrate, the obligatory activation of an enzyme precursor, the action of protease inhibitors, or a combination of these factors. By far the best characterized and perhaps most versatile proteolytic enzymes are the serine proteases. Together with their inhibitors, they regulate a great variety of physiological events. Whereas initially the different specificities of trypsin and chymotrypsin were exclusively ascribed to differences in the sequence and structure of the primary substrate-binding site (aspartic acid in trypsin vs. serine in chymotrypsin), this simple explanation had to be abandoned when Craik and coworkers (5) demonstrated that, in addition, two surface loops are changed, indicating that conformational changes at distant secondary binding sites are also required. It has also been shown that the introduction of a metal binding site by site-directed mutagenesis allows the interconversion of a protease belonging to the serine family into another that can be regulated like a zinc metallo protease (6, 7). However, the metal inhibits the serine protease but is essential for metalloprotease activity. A relative newcomer in the families of proteases are the caspases, which resemble each other in amino acid sequence, structure, and substrate specificity, as will be discussed in a paper to follow [G.S.Salvesen and V.M.Dixit (8)]. Another important recent advance is the isolation and characterization of proteasomes [R.Huber (9)]. One of the earliest and best understood cases of proteolytic processing is zymogen activation. It underlies a great variety of physiological regulations, particularly when coupled to consecutive activation reactions as in the cascades of blood

*To whom reprint requests should be addressed. E-mail: [email protected]. PNAS is available online at www.pnas.org.

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PROTEOLYTIC ENZYMES, PAST AND FUTURE

10963

coagulation, fibrinolysis, the complement reaction, and others. The key point here is that a signal can be specifically and irreversibly amplified every time a downstream inactive enzyme precursor is activated. Recent work, to be presented in the paper to follow (8), has demonstrated a specific role of the pro segment of activation, which early on was regarded a throwaway piece but in certain cases can act as an intramolecular inhibitor and as an intramolecular chaperon that assures proper folding of the active enzyme (10). Certain generalizations have emerged from these and related investigations. If I may borrow a page from Nancy Thornberry (11), most proteases are synthesized as inactive precursors (zymogens) that require limited proteolysis for activation. Because proteolysis is irreversible under physiological conditions, the generation of the uncleaved precursor requires de novo synthesis. All active proteases, including those that activate zymogens, are regulated by specific inhibitors. However, some protease precursors can regulate their own activation, e.g., trypsinogen, whereas others, e.g., plasminogen, do not require peptide bond cleavage for their activation. Proteolytic processing, like all proteolytic reactions, requires unique combinations of primary, secondary, and tertiary structures to permit interaction with substrate so as to form the reactive enzyme-substrate intermediate. Let me now make a major leap in time and discuss in brief how we reached the current era of research on proteolytic enzymes and what we can expect in the millennium that we are about to enter. Two major factors have expanded our conceptual horizons and endowed us with experimental tools of previously unimaginable powers of resolution. One factor is the application of the newly emerging concepts and methodologies of molecular and cell biology, such as DNA cloning and sequencing, site-directed mutagenesis, gene amplification, gene knockouts, phage display, and the wealth of information yielded by genomics research generally. The other major impetus came from a group of newly developed concepts and experimental approaches to the structure and function of proteins by mass spectroscopy (12), multidimensional NMR, and the use of computers for the prediction of protein structure based on various types of algorithms. To these one might add the methods of combinatorial chemistry as applied to proteins to scan and identify protein ligands of physiological significance. Although we are still far from understanding the rules of the in vivo folding of nascent polypeptide chains, the challenge lies in deriving the function of a protein from its known chemical and biological parameters and in learning how to design proteins of predetermined physiological properties. All of these developments, singly and in combination, expand our horizons and the goals that we are setting for their application to biology and medicine. The importance of proteolytic enzymes to the understanding of vital biological tasks is perhaps best illustrated by current trends in the study of viral proteases (13). In every known instance, the timing, placement, and mode of action of the virus encoded protease are somehow adapted to the conditions under which it operates within the viral environment. Two examples follow: in herpes viruses such as cytomegalovirus, the structure of the protease reveals a catalytic triad of His/His/Ser instead of the conventional Asp/His/Ser of the mammalian serine proteases and a single beta barrel structure per monomer instead of two in the mammalian serine proteases (13). Analogously, in adeno viruses the cysteine protease contains a Glu/His/Cys catalytic triad characteristic of cysteine proteases, but the seven alpha helices and a single five-stranded beta sheet are not seen in the parent protease (papain). In either case, the examples given demonstrate the ability of the virus proteases to adapt themselves to the evolution of functions within the limits of compatible protein structures (13). Other rapidly expanding areas of biological research involving well known proteases include those of apoptosis, the mediation of thrombin signaling by protease activated receptors, proteolytic processing in cholesterol metabolism, in the cell cycle, and the many others included in this issue of the Proceedings. It is no coincidence that industry and academia are almost equally represented in this audience, because intense cooperation between both is essential if we are to reap the full benefits of the advances and discoveries in both basic and applied research. 1. Northrop, J.H., Kunitz, M. & Herriott, R.M. (1938) Crystalline Enzymes (Columbia Univ. Press, New York). 2. Neurath, H. (1995) Protein Sci. 4, 1939–1943. 3. Barrett, A.J., Rawlings, N.D. & Woessner, J.F. (1998) in Handbook of Proteolytic Enzymes (Academic, New York), pp. xiii–xxix. 4. Kitamoto, Y., Yuan, X., Wu, Q., McCourt, D.W. & Sadler, J.E. (1994) Proc. Natl. Acad. Sci. USA 91, 7588–7592. 5. Perona, J.J. & Craik, C.S. (1995) Protein Sci. 4, 337–360. 6. Higaki, J.N., Fletterick, R.J. & Craik, C.S. (1992) Trends Biochem. Sci. 17, 100–104. 7. Klemba, M., Gardner, K.H., Marino, S., Clarke, N.D. & Regan, L. (1995) Struct. Biol. 2, 368–373. 8. Salvesen, G.S. & Dixit, V.M. (1999) Proc. Natl. Acad. Sci. USA 96, 10964–10967. 9. Groll, M., Heinemeyer, W., Jäger, S., Ullrich, T., Bochtler, M., Wolf, D.H. & Huber, R. (1999) Proc. Natl. Acad. Sci. USA 96, 10976–10983. 10. Cunningham, E.L., Jaswal, S.S. & Agard, D.A. (1999) Proc. Natl. Acad. Sci. USA 96, 11008–11014. 11. Thornberry, N.A. & Lazebnik, Y. (1998) Science 281, 1312–1316. 12. Cohen, S.L. (1996) Structure (London) 4, 1013–1016. 13. Babé, L.M. & Craik, C.S. (1997) Cell 91, 427–430.

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CASPASE ACTIVATION: THE INDUCED-PROXIMITY MODEL

10964

Caspase activation: The induced-proximity model

This paper was presented at the National Academy of Sciences colloquium “Proteolytic Processing and Physiological Regulation” held February 20–21, 1999, at the Arnold and Mabel Beckman Center in Irvine, CA. GUY S. SALVESEN*† AND VlSHVA M. DlXIT‡ *Programs in Cell Death and Aging Research, Burnham Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037; and ‡Department of Molecular Oncology, Genentech Inc., 460 Point San Bruno Boulevard, South San Francisco, CA 94080 ABSTRACT Members of the caspase family of proteases transmit the events that lead to apoptosis of animal cells. Distinct members of the family are involved in both the initiation and execution phases of cell death, with the initiator caspases being recruited to multicomponent signaling complexes. Initiation of apoptotic events depends on the ability of the signaling complexes to generate an active protease. The mechanism of activation of the caspases that constitute the different apoptosissignaling complexes can be explained by an unusual property of the caspase zymogens to autoprocess to an active form. This autoprocessing depends on intrinsic activity that resides in the zymogens of the initiator caspases. We review evidence for a hypothesis—the induced-proximity model—that describes how the first proteolytic signal is produced after adapter-mediated clustering of initiator caspase zymogens. Apoptosis is a mechanism that regulates cell number and is vital throughout the life of all animals. Though several different types of biochemical events have been recognized as important in apoptosis, perhaps the most fundamental is the participation of members of a family of cysteine-dependent, Asp-specific proteases known as the caspases (1–3). Caspases cleave a number of cellular proteins, and the process is one of limited proteolysis in which a small number of cuts, usually only one, are made in interdomain regions. Sometimes cleavage results in activation of the protein, sometimes in inactivation, but never in degradation, because their substrate specificity distinguishes the caspases as among the most restricted of endopeptidases. Singularly important in this context is that caspase zymogens are themselves substrates for caspases, such that some are able to activate others in a hierarchical relationship (Fig. 1). Thus, pathways exist to transmit signals via sequential caspase activations, and this event has been most extensively examined in apoptosis. It is relatively easy to imagine that the caspases operating at the bottom of the pathway are activated by the ones above. Until recently, the questions of how the first caspase in a pathway became activated and how the first death signal was generated were perplexing issues. Now, several groups have focused on this issue (4–7) and have arrived at a consensus to describe the intriguing operation of the initiation of the proteolytic pathways that execute apoptosis. Though the basic hypothesis is supported, many issues remain to be explained, not the least of which is the nature of the mechanism that governs the process. This paper reviews the support for the hypothesis—the induced-proximity model—and its current limitations. Apoptosis Triggered by Death Receptors. One of the most intensively studied pathways to cell death results from ligation of transmembrane death receptors belonging to the tumor necrosis factor-R1 (TNF-R1) family. After engagement by specific ligands, these receptors transmit a lethal signal that results in classic apoptotic cell death (8, 9). Because simple transfection of death receptors is usually sufficient to sensitize cells to a death ligand, it follows that the components required to transduce this signal reside in many cells. Thus TNF-R1 family members serve as a conduit for the transfer of death signals into the cell’s interior after interaction with their extracellular cognate ligands. The TNF-R1/TNF pair itself presents a rather complex pathway with which to dissect apoptosis initiation, because this receptor/ligand pair can signal either apoptosis or an antagonistic NF- B-mediated survival pathway, depending on the cellular context. The TNF-R1 homologue Fas (CD95/Apo-1) has been the paradigm of choice, because addition of its cognate ligand, FasL, or even receptor agonist antibodies rapidly signals cell death (10). Because agonist Fas antibodies can trigger apoptosis, it was possible to use them to isolate the components of the death-inducing signaling complex (DISC) that forms after Fas ligation (4, 11). A combination of yeast two-hybrid and proteinsequence analysis revealed a seemingly simple DISC, comprising Fas itself, the adapter molecule FADD, and caspase-8 (Fig. 1). This discovery revealed a potential solution to the perplexing problem of how the first proteolytic signal was generated during apoptosis, because it implicated a caspase directly in the triggering event. Before this work, receptors were thought to signal either by altering the phosphorylation status of key signaling molecules or by functioning as ion channels. Death receptors, such as Fas, signal by direct recruitment and activation of a protease (caspase-8). How exactly does the recruited zymogen become active? To understand this process as a basis for formulating an adequate hypothesis, one must understand the unusual properties of caspase zymogens that set them apart from most other proteases. Because, unlike most other proteases, simple expression of caspase zymogens in Escherichia coli usually results in their activation (12, 13). This activation results from processing that is a consequence of intrinsic proteolytic activity residing in the caspase zymogens. It is not caused by E.coli proteases, as indicated by the fact that catalytically disabled C285A (caspase-1 numbering convention) mutants fail to undergo processing. Self-Processing of Caspase Zymogens. In common with other protease zymogens (14), with notable exceptions (see Table 1), generation of an active caspase usually requires limited proteolysis (Fig. 2). The activating cleavage takes place within a short segment that, in the zymogen, connects the large and small subunits of the catalytic domain with both subunits containing essential components of the catalytic machinery. The location of cleavage within this segment need not be precise in vitro (15); nevertheless, the highly conserved Asp-

†To

whom reprint requests should be addressed. E-mail: [email protected]. PNAS is available online at www.pnas.org. Abbreviations: TNF, tumor necrosis factor; DISC, death-inducing signaling complex; DED, death-effector domain.

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297 (caspase-1 numbering convention) directs cleavage specificity within this segment in vivo. Proteolytic processing that results in activation usually occurs at this Asp residue, such that most activated caspases can process their own and other caspase zymogens, given sufficient time and a high enough concentration in vitro (16–18). The extent to which this processing occurs in vivo, however, is regulated by the residues surrounding Asp-297. For example, the sequence surrounding Asp-297 in the downstream executioner caspases 3 and 7 fits the extended substrate specificity of the initiator caspases 8 and 9 remarkably well (19). With the notable exception of at least caspase-2 (20), distinctions in substrate specificity within the caspase family fit closely to the S4–S1 subsite preferences deduced from synthetic peptidic substrates (19).

FIG. 1. The framework of apoptosis. Death may be signaled by direct ligand-enforced clustering of receptors at the cell surface, which leads to the activation of the “initiator” caspase-8 (casp-8). This caspase then directly activates the “executioner” caspases 3 and 7 (and possibly 6), which are predominantly responsible for the limited proteolysis that characterizes apoptotic dismantling of the cell. Alternatively, irreparable damage to the genome caused by mutagens, pharmaceuticals that inhibit DNA repair, or ionizing radiation leads to the activation of another initiator, caspase-9 (28). The latter event requires the recruitment of pro-caspase-9 to proteins such as Apaf-1, which requires the proapoptotic factor cytochrome c (cyto C) to be released from mitochondria (29). Though other modulators probably regulate the apoptotic pathway in a cell-specific manner (30), this framework is considered common to most mammalian cells. Table 1. Zymogenicities of some caspases compared with two serine proteases Zymogenicity Protease Caspase-3 >10,000 Caspase-8 100 Caspase-9 10 Trypsin >10,000 2–10 tPA Zymogenicity is defined as the ratio of the activity of a processed protease to the activity of the zymogen on any given substrate (27). Data for trypsin and tissue plasminogen activator (tPA) are taken from ref. 27. The interesting range of zymogenicity values displayed by members of the caspase family is mirrored by members of the chymotrypsin family, with trypsin and tPA shown for comparison. Presumably, enzmes such as tPA and caspase-9 have down played the requirement for proteolysis as a mechanism of substantially increasing their activities, because allosteric regulators substitute this function: fibrin for tPA and Apaf-1 for caspase-9. In the case of tPA, specific side-chain interactions, absent in other members of the chymotrypsin family, allow activity of the zymogen. However, in the absence of a molecular structure of the caspase-8 and caspase-9 zymogens, little evidence is available to explain the high activity of the unprocessed protein. One clue is suggested by the structure of active caspases 1 and 3, each of which is composed of two catalytic units thought to arise from the dimerization of monomeric zymogens (reiewed in ref. 3). If activation of zymogens of the initiator caspases-8, 9, and CED3 operates by clustering, then the clustering phenomenon may be explained by adapter-driven homodimerization of monomers. However, as detailed in Future Directions, the molecular mechanisms are far from clear.

The Induced-Proximity Hypothesis. Interestingly, depending on expression conditions, one can obtain either processed active caspase or unprocessed zymogen from the same construct, at least for caspases 3, 7, and 9 (15, 21, 22). For example, short induction times (3 hours) yield fully processed enzymes. Significantly, even very short expression times and low inducer concentrations have failed to yield caspase-8 zymogens in our studies (G.S. and H.Stennicke, unpublished work). Caspase-8 processes itself extremely rapidly on heterologous expression in E.coli, suggesting that the zymogen must possess significant intrinsic proteolytic activity, allowing for autoprocessing. These observation are the basis for the inducedproximity hypothesis for the operation of the DISC, the assembly of which forces a locally high concentration of caspase-8 zymogens in a process mediated by recruited FADD (Fig. 3). This clustering of zymogens possessing intrinsic enzymatic activity would allow for processing in trans as well as activation of the first protease in the cascade. The hypothesis would need to be tested by asking whether the zymogen form of caspase-8 possessed reasonable enzymatic activity. Because such a test could not be made by expressing the wild-type precursor, a nonprocessable mutant was generated by replacing the two Asp cleavage sites within the large/small subunit linker segment with Ala. These replacements enabled the generation of a “frozen” zymogen that could be obtained in quantity after expression in E. coli. Significantly, the frozen zymogen retained the same specificity against caspase inhibitors and synthetic substrates but cleaved these substrates at 1% of the rate of an equivalent concentration of fully processed enzyme. The mechanistic origin of this rate differential is currently unknown, but, significantly, the zymogenicity of caspase-8, the ratio of its activity as a fully active enzyme to the activity of its unprocessed zymogen, was 100 (4). The importance of zymogenicity is detailed in Table 1. Testing the Hypothesis. The in vitro observations on the high zymogenicity of caspase-8 suggested that a test of the induced autoprocessing hypothesis was mandated, preferably in vivo. With this mandate in mind, we generated a caspase-8 construct in which the DED domains of the zymogen were replaced by a myristoylation signal, followed by three tandem repeats of a derivative FK506 binding protein (FKBP). The latter had been designed by Schreiber and colleagues (23) to act as an artificial mimic of natural cellular recruitment processes. Artificial oligomerization of proteins carrying the FKBP domains was induced by treatment with the cell penetrant FK1012, a dimeric form of FK506. Ectopic expression of the catalytically active chimera was tolerated fairly well by two human cell lines, even in the presence of monomeric FK506. However, on addition of dimeric FK1012, the cells underwent apoptosis by a mechanism that depended on the catalytic function of the chimeric caspase-8, because replacing the catalytic Cys by Ser failed to elicit the same effect. This technique, later termed the “artificial death switch” (24), has taken a prominent position in the exploration of apoptosis initiation. These data, the in vitro observations on the zymogenicity of pro-caspase-8, and the artificially induced death of cells harboring the chimeric FKBP-caspase-8 are fully consistent with the induced-proximity model. Indeed, since this original description, the postmitochondrial initiator caspase-9 (7) and the Caenorhabditis elegans caspase CED3 (25) have both been implicated in congruent

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proximity activation mechanisms. Is this mechanism a common one for the basis of generating biochemical death signals? Possibly. However, a caveat must be added to the caspase-9 issue, because this caspase has a very low zymogenicity (22); in other words, it is almost as active before as it is after proteolytic processing! Thus, in the case of caspase-9, an alternative pathway may be used.

FIG. 2. Caspase activation by proteolysis. Caspases are synthesized as single-chain precursors that await activation within the cell. Activation usually proceeds in all caspases by cleavage at the conserved Asp-297 (caspase-1 numbering convention). After this activation, an as-yet undescribed conformational change is thought to occur, bringing the activity and specificity determinants (quarter circles in the linear precursor) into the correct alignment for catalysis. Frequently an N-terminal peptide is removed; however, the reason for this removal is obscure, because it is apparently not required for zymogen activation. In the example of caspase-8 shown in the figure, the N-peptide (sometimes called the prodomain) contains death-effector domains (DEDs) required for recruitment to the cytosolic face of death receptors. The crystal structures of caspases 1 and 3 reveal a dimer of small and large subunits in the active, processed state, and—it is assumed, though not specifically demonstrated—that this organization is the case for caspases in solution. The active sites in the putative dimer are shown as open circles. If the single-chain zymogens of caspases 8 and 9 are partly active, why are they not dangerous to healthy cells? They should cause a slow production of active executioner caspases. This question is most readily explained by the presence of endogenous caspase inhibitors, members of the IAP (inhibitor of apoptosis protein) family (31). Members of this family inhibit executioner caspases 3 and 7, and we propose that they present a barrier to caspase activity that must be exceeded before sufficient execution potential can be achieved. Thus, in the presence of IAPs, a little caspase activation is acceptable, because it would be rapidly saturated by the inhibitors. It is only when a sufficient concentration of activated executioner caspases builds up that apoptosis occurs. In this hypothesis, the IAPs regulate the apoptotic threshold.

FIG. 3. Model for the operation of the DISC. Assembly of the DISC occurs in a hierarchical manner. On ligation of Fas, its “death domain” (white circle) binds to a homologous domain in the adapter FADD, which in turn recruits the zymogen of caspase-8 by a homophilic interaction requiring the homologous DEDs (black circles). Immediately after recruitment, the zymogen is processed by an adjacent zymogen, resulting in proteolytic activation and origination of active caspase-8 as the initiating death signal. Activation is thought to result from cleavage at Asp-297 (caspase-1 numbering convention). Presumably, the active form of caspase-8 (designated as a dimer as seen in the structures of active caspases 1 and 3) releases itself from the adapter after proteolytic removal of the N-terminal DED, though it is not clear how the endogenous activated enzyme distributes in the cell. Future Directions. Notwithstanding the attractiveness of the induced-proximity model, there remain a number of open questions. For example, although the data support the hypothesis, the molecular mechanisms of the event(s) have not been explained, and there are a number of issues that need to be addressed in the near future. These issues are as follows. (i) Must the processed caspase-8 be released from the DISC to diffuse toward its downstream substrates? (ii) Does activation require dimerization, a consensus for the catalytic form of caspases 1 and 3 at least? (iii) Does processing occur in cis (intramolecular) or in trans (intermolecular)? (iv) Must the zymogens be specifically aligned within the recruitment complex, and how many zymogen molecules constitute an activation locus? (v) Is the minimal operative DISC as simple as the one depicted in Fig. 3, or are other proteins required (26)? These questions cut to the heart of uncertainties surrounding the fundamental activation mechanism of all the caspases, and each is (in principle) answerable by generating specific mutants and by using the artificial death-switch technique. Perhaps it is already possible to settle the issue of cis versus trans processing; in our hands, it is rarely possible to observe activation of caspase zymogens in the nanomolar range, but on artificial concentration toward the micromolar range, one observes processing and activation. This observation would imply a secondorder reaction, which is most easily understood in terms of trans processing. Indeed, this proposal makes sense, because it is much easier to regulate zymogen activation in trans than in cis. The answers to these questions will require the molecular structure of at least one caspase zymogen (preferably caspase-8). Their resolution will certainly lead to a better understanding of the molecular mechanism of the DISC, with the attendant possibilities of interfering therapeutically to either initiate or prevent the commitment step in deathreceptor-mediated apoptosis. This work was supported by grants from the National Heart, Lung, and Blood Institute, National Institute on Aging, and National Institute of Neurological Disorders and Stroke.

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1. Salvesen, G.S. & Dixit, V.M. (1997) Cell 91, 443–446. 2. Cohen, G.M. (1997) Biochem. J. 326, 1–16. 3. Thornberry, N.A. & Lazebnik, Y. (1998) Science 281, 1312–1316. 4. Muzio, M., Stockwell, B.R., Stennicke, H.R., Salvesen, G.S. & Dixit, V.M. (1998) J. Biol. Chem. 273, 2926–2930. 5. Martin, D.A., Siegel, R.M., Zheng, L. & Lenardo, M.J. (1998) J. Biol. Chem. 273, 4345–4349. 6. Yang, X., Chang, H.Y. & Baltimore, D. (1998) Mol. Cell 1, 319–325. 7. Srinivasula, S.M., Ahmad, M., Fernandes-Alnemri, T. & Alnemri, E.S. (1998) Mol. Cell 1, 949–957. 8. Ashkenazi, A. & Dixit, V.M. (1998) Science 281, 1305–1308. 9. Ware, C.F., Santee, S. & Glass, A. (1998) in The Cytokine Handbook (Academic, London), 3rd Ed., pp. 549–592. 10. Nagata, S. & Goldstein, P. (1995) Science 267, 1449–1456. 11. Boldin, M.P., Goncharov, T.M., Goltsev, Y.V. & Wallach, D. (1996) Cell 85, 803–815. 12. Orth, K., O’Rourke, K., Salvesen, G.S. & Dixit, V.M. (1996) J. Biol Chem. 271, 20977–20980. 13. Stennicke, H.R. & Salvesen, G.S. (1997) J. Biol. Chem. 272, 25719–25723. 14. Neurath, H. (1989) Trends Biochem. Sci. 14, 268–271. 15. Zhou, Q. & Salvesen, G.S. (1997) Biochem. J. 324, 361–364. 16. Srinivasula, S.M., Ahmad, M., Fernandes-Alnemri, T., Litwack, G. & Alnemri, E.S. (1996) Proc. Natl. Acad. Sci. USA 93, 14486–14491. 17. Muzio, M., Salvesen, G.S. & Dixit, V.M. (1997) J. Biol. Chem. 272, 2952–2956. 18. Slee, E.A., Harte, M.T., Kluck, R.M., Wolf, B.B., Casiano, C.A., Newmeyer, D.D., Wang, H.G., Reed, J.C., Nicholson, D.W., Alnemri, E.S., et al. (1999) J. Cell Biol. 144, 281–292. 19. Thornberry, N.A., Rano, T.A., Peterson, E.P., Rasper, D.M., Timkey, T., Garcia-Calvo, M., Houtzager, V.M., Nordstrom, P.A., Roy, S., Vaillancourt, J.P., et al. (1997) J. Biol. Chem. 272, 17907–17911. 20. Talanian, R.V., Quinlan, C., Trautz, S., Hackett, M.C., Mankovich, J.A., Banach, D., Ghayur, T., Brady, K.D. & Wong, W.W. (1997) J. Biol Chem. 272, 9677–9682. 21. Stennicke, H.R., Jurgensmeier, J.M., Shin, H., Deveraux, Q., Wolf, B.B., Yang, X., Zhou, Q., Ellerby, H.M., Ellerby, L.M., Bredesen, D., et al. (1998) J. Biol. Chem. 273, 27084–27090. 22. Stennicke, H.R., Deveraux, Q.L., Humke, E.W., Reed, J.C., Dixit, V.M. & Salvesen, G.S. (1999) J. Biol Chem. 274, 8359–8362. 23. Spencer, D.M., Belshaw, P.J., Chen, L., Ho, S.N., Randazzo, F., Crabtree, G.R. & Schreiber, S.L. (1996) Curr. Biol. 6, 839–847. 24. MacCorkle, R.A., Freeman, K.W. & Spencer, D.M. (1998) Proc. Natl. Acad. Sci. USA 95, 3655–3660. 25. Yang, X., Chang, H.Y. & Baltimore, D. (1998) Science 281, 1355–1357. 26. Imai, Y., Kinura, T., Murakami, A., Yajima, N,, Sakamaki, K. & Yonehara, S. (1999) Nature (London) 398, 777–785. 27. Tachias, K. & Madison, E.L. (1996) J. Biol. Chem. 271, 28749– 28752. 28. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S.M., Ahmad, M., Alnemri, E.S. & Wang, X. (1997) Cell 91, 479–489. 29. Zou, H., Henzel, W.J., Liu, X., Lutschg, A. & Wang, X. (1997) Cell 90, 405–413. 30. Green, D.R. & Reed, J.C. (1998) Science 281, 1309–1312. 31. Deveraux, Q., Takahashi, R., Salvesen, G.S. & Reed, J.C. (1997) Nature (London) 388, 300–304.

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STRUCTURAL ASPECTS OF ACTIVATION PATHWAYS OF ASPARTIC PROTEASE ZYMOGENS AND VIRAL 3C PROTEASE PRECURSORS

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Structural aspects of activation pathways of aspartic protease zymogens and viral 3C protease precursors

This paper was presented at the National Academy of Sciences colloquium “Proteolytic Processing and Physiological Regulation” held February 20–21, 1999, at the Arnold and Mabel Beckman Center in Irvine, CA. AMIR R. KHAN*, NINA KHAZANOVICH-BERNSTEIN, ERNST M. BERGMANN, AND MICHAEL N. G. JAMES† Medical Research Council Group in Protein Structure and Function, Department of Biochemistry, University of Alberta, Edmonton, Alberta T6G 2H7, Canada ABSTRACT The three-dimensional structures of the inactive protein precursors (zymogens) of the serine, cysteine, aspartic, and metalloprotease classes of proteolytic enzymes are known. Comparisons of these structures with those of the mature, active proteases reveal that, in general, the preformed, active conformations of the residues involved in catalysis are rendered sterically inaccessible to substrates by the residues of the zymogens’ N-terminal extensions or prosegments. The prosegments interact in nonsubstrate-like fashions with the residues of the active sites in most of the cases. The gastric aspartic proteases have a well-characterized zymogen conversion pathway. Structures of human progastricsin, the inactive intermediate 2, and active human pepsin are known and have been used to define the conversion pathway. The structure of the zymogen precursor of plasmepsin II, the malarial aspartic protease, shows a new twist on the mode of inactivation used by the gastric zymogens. The prosegment of proplasmepsin disrupts the active conformation of the two catalytic aspartic acid residues by inducing a major reorientation of the two domains of the mature protease. The picornaviral 2A and 3C proteases have a chymotrypsin-like tertiary structure but with a cysteine nucleophile. These enzymes cleave themselves from the viral polyprotein in cis (intramolecular cleavage) and carry out trans cleavages of other scissile peptides important for the virus life cycle. Although the structure of the precursor viral polyprotein is unknown, it probably resembles the organization of the proenzymes of the bacterial serine proteases, subtilisin, and α-lytic protease. Cleavage of the prosegment is known to occur in cis for these precursor molecules. Zymogens of proteolytic enzymes consist of the intact protease with an N-terminal extension. Conversion of the inactive zymogen to the mature, active protease requires limited proteolysis usually of a single peptide bond (1). Molecular rearrangements accompany the proteolytic removal of the prosegment of the zymogen, eventually leading to the mature protease. The prosegments of the zymogens range in size from two residues for some of the granzymes to more than 150 residues for a-lytic protease, a bacterial serine protease (2). The conversion of zymogens to the respective active enzymes is achieved by several different mechanisms (3). The active serine proteases of the chymotrypsin family result from limited proteolysis of the zymogens by convertases. For example, the cascade of the blood-clotting enzymes (4) involves the conversion of inactive forms (e.g., prothrombin) to active forms of the enzyme (thrombin) by a highly specific catalytic cleavage by another of the clotting enzymes (factor Xa). On the other hand, simply changing the pH of the solution in which the gastric aspartic protease zymogens are dissolved from
6.5 to 3.0 (an increase in [H+] of
3,100-fold) is sufficient to bring about the conversion (5). In a similar fashion, the zymogens of the papain-like cysteine proteases are converted to the active enzymes in a pH-regulated fashion. The in vitro activation of propapain is consistent with an initial intramolecular cleavage event (6). The conversion of procarboxypeptidase is initiated by trypsin cleavage of the Arg-99p-Ala-1 bond at the prosegment to mature enzyme junction (7). Prostromelysin-1 can be converted to the active form by other proteolytic enzymes, heat, or the presence of organomercurial agents (8). There are some generalities regarding zymogen conversion that one can make in light of the three-dimensional structures of both the zymogens and the respective active enzymes (3). First, the residues that constitute the active sites of the protease portions of the zymogens have virtually identical conformations to those of the mature, active proteases. The major exceptions are the serine proteases of the chymotrypsin family. The activation process involves the formation of an ion pair between the newly formed N-terminal residue Ile-16 NH3+ and the β-carboxylate of Asp-194 (9), which triggers the conformational changes that form the oxyanion binding pocket and the active conformation of the S1 specificity pocket [the nomenclature of Schechter and Berger (10) is used throughout this manuscript]. Second, the preformed active sites of the protease portions of zymogens are generally not accessible to substrates because residues of the prosegments sterically block the approach to the active sites. This statement does not hold for the chymotrypsin-like serine proteases as the active sites of these zymogens are able to bind protein inhibitors that induce conformational changes that form the oxyanion hole in spite of the ion pair involving Asp-194 and Ile-16 being absent (11). Proteolysis of the portion of the prosegments that interact with the active site residues is prevented in several different ways. In prostromelysin the prosegment passes through the active site in the reverse polypeptide direction (N→C) relative to substrates or transition state mimics (12). A reverse orientation of the prosegment blocking the active site in the cysteine protease zymogens also has been observed in the structures of rat procathepsin B (13) and human procathepsin L (14). The region of the prosegment of the gastric aspartic proteases interacting with the catalytic residues Asp-32 and Asp-215 (porcine pepsin numbering) most intimately, includes a highly conserved lysine at position 36p (the residue numbers of the prosegment are followed by p). The εNH3+ group of Lys-36p forms an ion pair with each of the two active site carboxylate groups (15).

Conversion of Gastric Aspartic Protease Zymogens The molecular structures of human progastricsin (16), activation intermediate 2 of human gastricsin (17), and a structural

*Present address: Department of Molecular and Cellular Biology, Harvard University, 7 Divinity Avenue, Cambridge, MA 02138. †To whom reprint requests should be addressed. E-mail: [email protected]. PNAS is available online at www.pnas.org. Abbreviation: HPV, human polio virus.

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homolog to mature, human gastricsin, human pepsin 3 (18) allow one to construct a reasonably detailed view of the pathway followed in the conversion of the inactive zymogen to the active protease. Fig. 1 shows stereo ribbon diagrams of each of these three molecular structures. Fig. 2 shows a diagrammatic view of the conversion pathway. This pathway is a general pathway for the gastric aspartic proteases but the individual enzymes differ in detail.

FIG. 1. Structures on the conversion pathway of the aspartic protease zymogen progastricsin. The structure of human gastricsin is not known; the human pepsin structure therefore has been used as a model for gastricsin. This figure, as well as Figs. 3–6 have been prepared with BOBSCRIPT (19) and RASTER 3D (20). (A) The structure of human progastricsin (16) represented in stereo. The residues of the prosegment (Ala-1p to Leu-43p) are in green, those of the gastricsin portion of the zymogen are in blue except for those regions that undergo large conformational changes, Ser-1 to Ala-13, Phe-71 to Thr-81 and Tyr-125 to Ala-136, which are represented in mauve. The promature junction is Leu-43p-Ser-1, the peptide bond cleaved intramolecularly is Phe-26p to Leu-27p. The side chains of Asp-32 and Asp-217 are represented in red. (B) Stereo view of the molecular structure of intermediate 2 on the activation pathway of human gastricsin (17). The color scheme used is the same as in A. The residues missing on this figure, Leu-22p to Phe-26p and Ser-1, are disordered in the structure, and there is no interpret able electron density for them on the maps. The water molecule bound between the two carboxyl groups of Asp-32 and Asp-217 is shown as a red sphere. The final step in the conversion involves the dissociation of the peptide Ala-1p to Phe-26p from gastricsin with the N-terminal residues of gastricsin, Ser-1 (N-ter) to Ala-13, replacing the N-terminal β-strand of the prosegment. (C) The structure of human pepsin (18) shown as a model of human gastricsin. The regions of gastricsin that undergo large conformational changes from their positions in progastricsin are shown in pink, and the active site aspartates with the bound catalytic H2O molecule are colored red. Reproduced with permission from ref. 3.

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FIG. 2. A diagrammatic representation of the conversion pathway of progastricsin to gastricsin. The prosegment of progastricsin (I), A1p to L43p, has three helical segments and a net positive charge forming several ion pair interactions and electrostatic stabilization with the mature portion of the zymogen. The highly conserved K37p interacts directly with the catalytic aspartates Asp-32 and Asp-217. Lowering the pH of the solution below 4.0 converts progastricsin into intermediate 1 (26) represented by II. Refolding of the prosegment in the vicinity of the active gastricsin brings the first scissile bond Phe-26p-Leu-27p to the exposed active site aspartates (III). Cleavage at the premature junction, Leu-43p-Ser-1, (IV) is likely an intermolecular cleavage (28) and results in intermediate 2 (V), a molecular species that consists of Ala-1p to Phe-26p noncovalently associated with mature gastricsin (17). The final step in the conversion results from the dissociation of the Nterminal peptide 1–26 of the prosegment and refolding the residues Ser-1 to Ala-13 to replace the region of the prosegment in the six-stranded β-sheet of gastricsin (VI). Progastricsin consists of a single polypeptide chain of 372 aa (21). The N-terminal extension or prosegment is 43 aa in length and comprises residues Ala-1p to Leu-43p. The prosegment is folded into a compact domain having an initial extended β-strand (Val-3p to Lys-8p) followed by three helical segments: Ile-13p to Lys-20p, Leu-22p to Arg-28p, Pro-34p to Arg-39p (16). The third helical segment (a 310 helix) packs against the active site residues and the εNH3+ group of a conserved lysine residue (Lys-37p in progastricsin) forms ion pair interactions with the carboxyl groups of the two catalytic aspartates, Asp-32 and Asp-217. Two tyrosine side chains Tyr-38p and Tyr-9 form symmetric H-bonded interactions with the carboxylates of Asp-217 and Asp-32, respectively, further restricting access to the active site. The phenolic side chain of Tyr-9 occupies the S1 binding pocket; Tyr-38p is in the S1 binding pocket. The tertiary structure of the prosegment (Leu-1p to Tyr-37p) in porcine pepsinogen (15) is virtually identical to that described above for progastricsin (16). The polypeptide chain from Tyr-38p to Tyr-9 in progastricsin adopts a conformation that is different from the equivalent segment of chain in the pepsinogens (15, 22). As well, a portion of the polypeptide chain in gastricsin Tyr-125 to Ala-136 (Fig. 1A) is displaced from the position that this chain segment occupies in all other aspartic protease zymogens and active enzymes (16). The trigger for initiating the conversion of the gastric aspartic protease zymogens is a drop in pH (5). At neutral pH, the structures of the zymogens are stabilized by the electrostatic interactions of the ion pairs and the inactive conformation is maintained (16). However, when the zymogens reach the acid pH (
2.0) of the lumen of the stomach, the carboxylate groups become protonated and the repulsive interactions among the net positive charges of the prosegment destabilize its interactions in the active site of the protease. Kinetic studies in the late 1930s showed that the conversion of porcine pepsinogen into pepsin was an autocatalytic process (5, 23). In addition, the fact that the loss of pepsinogen was not accompanied by an equivalent increase in the appearance of pepsin implied the presence of intermediate species on the pathway (5). Spectroscopic studies of this conversion process established that there are conformational changes (24) in the 5-ms to 2-s time scale (25). Rapidly changing the pH back to neutrality can reverse these conformational changes. Biochemical studies of the conversion of human progastricsin to gastricsin showed the presence of at least two intermediates (26). Intermediate 1 is the species formed rapidly after the pH was dropped below 4.0. The prosegment is unfolded in intermediate 1 and the active site of gastricsin is exposed and accessible to substrates. The first hydrolytic event detected during the activation of progastricsin is the intramolecular cleavage of the Phe-26p to Leu-27p peptide bond (26). Subsequently, an intermolecular cleavage at the Leu-43pSer-1 peptide bond (the promature junction) results in the formation of transient intermediate 2 that can be stabilized by transferring the pH to neutrality (>6.5). The resulting molecular species has been characterized biochemically (26) and comprises residues Ala-1p to Phe-26p noncovalently associated with mature gastricsin (Ser-1 to Ala-329). Intermediate 2 recently has been characterized structurally (17), and its structure is depicted in Fig. 1B. The β-strand (Val-3p to Lys-8p) is in the same position as observed in the structure of progastricsin. In addition, the first helix (Ile-13p to Lys-20p) is intact and is very similarly oriented as it is in the zymogen structure. The two catalytic aspartates, Asp-32 and Asp-217, have a water molecule bound between them in the same position as the nucleophilic water observed in the native structures of all mature aspartic proteases (27). The S1 binding site is occluded, however, as the side chain of Tyr-9 still forms a hydrogen bond with the carboxylate of Asp-32. The segment Tyr-125 to Ala-136 has moved from its position in progastricsin

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STRUCTURAL ASPECTS OF ACTIVATION PATHWAYS OF ASPARTIC PROTEASE ZYMOGENS AND VIRAL 3C PROTEASE PRECURSORS

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(Fig. 1A) to the position and conformation common among the mature enzymes whose structures have been solved (Fig. 1 B and C). The ion pair Arg-14p to Asp-11 in intermediate 2 at pH
6.5 stabilizes the N-terminal peptide, Ala-1p to Phe-26p, in its original location in the zymogen preventing the N-terminal residues of gastricsin (Ser-1 to Ala-12) from adopting their final position in the mature enzyme. On the other hand, intermediate 2 would be relatively short-lived at acid pH values